{'fraction_non_alphanumeric': 0.030638410362602987, 'fraction_numerical': 0.023624267524342425, 'mean_word_length': 5.044205368555603, 'pattern_counts': {'":': 0, '<': 5, '<?xml version=': 0, '>': 1, 'https://': 0, 'lorem ipsum': 0, 'www.': 0, 'xml': 0}, 'pii_count': 0, 'substrings_counts': 0, 'word_list_counts': {'cursed_substrings.json': 0, 'profanity_word_list.json': 0, 'sexual_word_list.json': 0, 'zh_pornsignals.json': 0}}
EASTERN OYSTER LARVAL TRANSCRIPTOMES IN RESPONSE TO PROBIOTIC AND PATHOGENIC BACTERIA

Oysters are described as keystone species serving an important ecological role. As filter-feeders they help in maintaining water quality. Oyster reefs provide refuge and support to different organisms. The eastern oyster, Crassostrea virginica, native to the East Coast of United States and Gulf of Mexico is a part of the rapidly growing aquaculture industry. Aquaculture production depends on a healthy and constant supply of oyster larvae that are provided by hatcheries. Several hatcheries on the east coast that provide C. virginica seed to oyster farms face significant losses owing to Vibrio infections causing massive larval mortalities. Use of antibiotics is avoided due to possibility of development of antibiotic resistance. The probiotic bacteria, Phaeobacter inhibens S4 and Bacillus pumilus RI06-95 have been shown to successfully protect C. virginica larvae from V. coralliilyticus RE22 infection. Use of these probiotics in hatcheries can reduce mortalities due to disease thereby avoiding significant economic losses. In order to design best practices for probiotic use it is crucial to understand their mechanisms of action. There has been great progress in understanding the components of oyster immune system, its functioning in response to various stimuli and its uniqueness as compared to other organisms. This is in part due to availability of sophisticated tools like high throughput sequencing and various –omics analyses such as proteomics, genomics and transcriptomics and partly due to interest in controlling diseases affecting aquaculture. As such most of our knowledge is based on studies that focus on oyster-pathogen or oyster-environmental stimuli interaction. Little is known about the effect of bacteria other than pathogens on the oysters. Moreover, very little about larval immunity of eastern oyster, C. virginica. This is the first study to investigate the effect of both pathogen and probiotic bacteria on C. virginica larval immunity using transcriptomes. The aim of this study is to test the safety and efficacy of formulated probiotic Bacillus pumilus RI06-95 in a hatchery, understand the mechanisms of action of both probiotics and to characterize the effect of V. coralliilyticus RE22 infection on the larval immune system of eastern oysters. Chapter 1 reviews the current knowledge of oyster immune system and the mechanisms of action of probiotics especially mechanisms related to immunomodulation of innate immunity. Previous studies have demonstrated successful protection of C. virginica larvae from V. coralliilyticus RE22 infection in a laboratory based setting as well as in a hatchery using laboratory grown cultures of probiotics. The ultimate use of the probiotics is in a hatchery setting, which would require easy to use and stable formulation of the probiotics instead of time consuming laboratory-grown probiotic cultures that are viable for only a short duration of time. Chapter 2 discusses methods of formulation of probiotic Bacillus pumilus RI0695, testing the formulation in a hatchery and its effect on larval survival at the hatchery and post V. coralliilyticus RE22 experimental challenge in the laboratory. A spray dried formulation of Bacillus pumilus RI06-95 was both shelf-stable and effective in protecting C. virginica larvae from V. coralliilyticus RE22 challenge. The formulation did not show any adverse effects on the larvae during the course of the trial. Chapter 3 investigates the host–pathogen interaction between C. virginica larvae and V. coralliilyticus RE22 using transcriptomes produced after experimental challenge. Exposure of larvae to the pathogen for 6 hours provided information of the changes in the larval oysterimmune system brought about by the pathogen in the early stages of disease. Overall, despite upregulation of several pattern recognition receptors, immune signaling pathways leading to the production of antimicrobial effectors, such as protease inhibitors and the pore forming protein perforin-2, were suppressed by V. coralliilyticus RE22. The transcriptomic evidence suggests that lack of an adequate immune response to thwart the infection of RE22, combined with a high metabolic load and decreased feeding, leads to large-scale mortalities of C. virginica larvae. This research allows for a better understanding of the disease process caused by V. coralliilyticus RE22 in larval eastern oysters. Chapter 4 investigates the effect of exposure to non-pathogenic probiotic bacteria P. inhibens S4 and B. pumilus RI06-95 on the immune system of the host, C. virginica larvae. It presents evidence of immunomodulation of C. virginica larval immunity by both probiotic organisms. High upregulation of immune effectors such as serine protease inhibitors is seen in larval oysters after short exposures to the probiotic (6 and 24h) in the laboratory as well as after exposure for several days during a hatchery trial. Other important modulations that help larvae protect themselves from V. coralliilyticus RE22 infection include activation of pathogen receptors and signaling pathways, modulation of mucin genes, and upregulation of pore-forming protein perforin-2. Chapter 5 summarizes and advocates the of use of probiotics in the larviculture of C. virginica and suggests their potential role in limiting vibriosis.

the effect of both pathogen and probiotic bacteria on C. virginica larval immunity using transcriptomes. The aim of this study is to test the safety and efficacy of formulated probiotic Bacillus pumilus RI06-95 in a hatchery, understand the mechanisms of action of both probiotics and to characterize the effect of V. coralliilyticus RE22 infection on the larval immune system of eastern oysters.
Chapter 1 reviews the current knowledge of oyster immune system and the mechanisms of action of probiotics especially mechanisms related to immunomodulation of innate immunity. Previous studies have demonstrated successful protection of C. virginica larvae from V. coralliilyticus RE22 infection in a laboratory based setting as well as in a hatchery using laboratory grown cultures of probiotics. The ultimate use of the probiotics is in a hatchery setting, which would require easy to use and stable formulation of the probiotics instead of time consuming laboratory-grown probiotic cultures that are viable for only a short duration of time.
Chapter 2 discusses methods of formulation of probiotic Bacillus pumilus RI06-95, testing the formulation in a hatchery and its effect on larval survival at the hatchery and post V. coralliilyticus RE22 experimental challenge in the laboratory. A spray dried formulation of Bacillus pumilus RI06-95 was both shelf-stable and effective in protecting C. virginica larvae from V. coralliilyticus RE22 challenge. The formulation did not show any adverse effects on the larvae during the course of the trial.
Chapter 3 investigates the host-pathogen interaction between C. virginica larvae and V. coralliilyticus RE22 using transcriptomes produced after experimental challenge.
Exposure of larvae to the pathogen for 6 hours provided information of the changes in the larval oysterimmune system brought about by the pathogen in the early stages of disease. Overall, despite upregulation of several pattern recognition receptors, immune signaling pathways leading to the production of antimicrobial effectors, such as protease inhibitors and the pore forming protein perforin-2, were suppressed by V. coralliilyticus RE22. The transcriptomic evidence suggests that lack of an adequate immune response to thwart the infection of RE22, combined with a high metabolic load and decreased feeding, leads to large-scale mortalities of C. virginica larvae. This research allows for a better understanding of the disease process caused by V. coralliilyticus RE22 in larval eastern oysters.
Chapter 4 investigates the effect of exposure to non-pathogenic probiotic bacteria P. inhibens S4 and B. pumilus RI06-95 on the immune system of the host, C. virginica larvae. It presents evidence of immunomodulation of C. virginica larval immunity by both probiotic organisms. High upregulation of immune effectors such as serine protease inhibitors is seen in larval oysters after short exposures to the probiotic (6 and 24h) in the laboratory as well as after exposure for several days during a hatchery trial. Other important modulations that help larvae protect themselves from V. coralliilyticus RE22 infection include activation of pathogen receptors and signaling pathways, modulation of mucin genes, and upregulation of pore-forming protein perforin-2.
Chapter 5 summarizes and advocates the of use of probiotics in the larviculture of C. virginica and suggests their potential role in limiting vibriosis.
v ACKNOWLEDGEMENTS I want to take this opportunity to sincerely thank my advisor Dr. Marta Gomez-Chiarri, who kept a sense of humor even when I sometimes lost mine. She is a role model for an excellent advisor, a female scientist and the kindest and warmest boss I have ever met. She welcomed me to join her lab in the middle of my program ensuring a very smooth transition. She encouraged free thought in my research, while occasionally giving me a nudge in the right direction. In her company, I always found myself working the hardest due to the confidence she entrusted in me. I am so thankful and lucky to have her as my mentor.
I also want to thank Dr. Bethany Jenkins who championed me during my transition and pivoted my research career by helping me pick the right lab. I want to express my sincere thanks to all the hatchery managers that I hounded every spring for sending me oyster larvae and who patiently shipped millions of these critters for free to be sacrificed in the name of science. Special mention to Mike Congrove, Lauren Shirley, x

Immunity in oysters
Oysters are sessile filter feeding animals that provide important ecological and economical services. As such immunological studies to understand disease resistance and improve aquaculture practices has given a boost in our understanding of the oyster immunology. Although some research suggests presence of immunological memory (Green et al., 2015) it is generally recognized that oysters lack adaptive immunity and only possess innate immunity. The circulating phagocytic hemocytes form the cellular branch of the innate immunity in oysters. The production of antimicrobial effectors via activated signaling pathways due to recognition of PAMPs (pathogen-associated molecular patterns) by PRRs (pattern recognition receptors) forms the humoral branch of innate immunity (Wang et al., 2018). Current research of the oyster immunity is reviewed below with emphasis on (i) Recognition (ii) Signaling pathways (iii) Effectors
Several of these PRRs are highly diversified in oysters (Zhang et al., 2014, Zhang et al., 2015. C-type lectins require a special mention since they are not only involved in pathogen recognition but also in activation of complement cascade.

Signaling pathways
Signals transmitted by receptors allow activation of several signaling pathways like TLR signaling pathway, NF-kB signaling pathway, mitogen-activated protein kinase (MAPK) signaling cascade, prophenol/phenol oxidase cascade and complement pathway in oysters. Sophisticated tools like whole genome sequencing and -omics analysis have led to tremendous progress in understanding molecules involved in these pathways that are common with other organisms as well unique to oysters. The TLR/NF-kB signaling pathway is a crucial pathway that upon recognition by TLR receptors activate transcription factors facilitating production of effectors like cytokines, interleukins, antimicrobial peptides (AMPs) and others (Gerdol et al., 2018).
MyD88 serves as a critical cytosolic adaptor modulating TLR signaling pathway and Pacific oyster genome encodes an expanded set of 10 MyD88 genes (Zhang et al., 2015).
MAPK pathway comprises of many protein kinases and its active involvement in oyster immunity is evidenced by their activation upon bacterial exposures (Qu et al., 2016).
Although studies support existence of a complement pathway in bivalves (Gerdol et al., 2015, Li et al., 2015 the exact components and mechanisms of activation remain to be identified (Gerdol et al., 2018).

Effectors
Broad ranged effectors are produced upon induction of signaling pathways by PRR recognition and function in elimination of pathogens. These include antimicrobial peptides (AMPs), defensins, lysozymes, cytokines, protease inhibitors, antioxidant enzymes and acute phase proteins. Serine protease inhibitors have been identified for their role as important effectors in granting resistance to pathogens (La Peyre et al. 2010, Xue et al. 2006. Enzymes, such as superoxide dismutase, catalase and glutathione peroxidase defend oysters by eliminating reactive oxygen species (ROS). They are important especially during increased oxidative stress caused by pathogen infection. Another important member of effectors are the heat shock proteins (HSPs) that help oysters modulate stress response and protect them from environmentally induced cellular damage caused by a variety of stressors (Wang et al., 2018).

Apoptosis and autophagy
Apoptosis, programmed cell death is an extremely important process in oysters involved in immune system homeostasis and function, defense against parasite and pathogens and self/non-self recognition. The baseline apoptosis rates observed in circulating and resident hemocytes in oysters is high (Sokolova, 2009). Apoptosis in oysters has two major pathways intrinsic and extrinsic. The main players consist of caspases and inhibitors of apoptosis (IAPs) that regulate the process. Apoptosis limits the spread of pathogen while preventing inflammatory damage of surrounding tissues (Sokolova, 2009). Although apoptosis has been studied for a long time the exact functional relevance of its modulation by biotic and abiotic factors is still unknown in bivalves (Gerdol et al., 2018, Wang et al., 2018.
Autophagy plays a housekeeping role in organisms and is important in innate immunity.
It is activated in oysters in response to bacterial, viral and environmental stimuli (Gerdol et al., 2018, Wang et al., 2018. Its role in protecting Pacific oysters from viral and bacterial challenge was demonstrated recently (Moreau et al., 2015) but a lot more remains to be investigated.

Mucosal immunity
Mucus forms an external barrier of defense and plays a key role in host-microbe interactions. Mucus consisting of crosslinked glycoproteins forms a physical barrier to microbes and contains a myriad of effectors that defend the host from infection . These include enzymes like lysozymes, hydrolases and proteases, AMPs, antioxidants and lectins to name a few . Mucus composition can affect pathogen adhesion and production of components is often regulated by them (Linden et al., 2008, Allam and. This understudied topic is a crucial part of the innate immunity in oysters and needs further exploration.
Most of the knowledge of oyster immunity is based on a large body of research that is centered on bacterial and viral pathogens and environmental stressors but we know very little about the impact of friendly or beneficial bacteria on the immune system of oysters.
Addressing this dearth of knowledge might reveal important novel insights in the oyster immune system. The next section of this review discusses the effect of probiotics on different organisms focusing especially on their impact on immune system.  (Vine et al., 2006, Ray et al. 2012. Increasing nutrient availability and stimulation of growth through increased volatile fatty acids production by probiotics has been studied in poultry industry as well (Ajuwon., 2016).

Production of inhibitory compounds
Probiotics produce or stimulate production of several non-specific compounds that are effective in inhibiting pathogen growth including, antimicrobial compounds (hydrogen peroxide, nitric acid and bacteriocins), siderophores, proteases and lysozymes. A nonpathogenic strain Vibrio mediterranei 1 produces bacteriocin-like inhibitory substance against Vibrio parahaemolyticus spp (Carraturo et al., 2006). In fact, bacteriocin production allows probiotics to compete within complex microbial communities and influence the health of the host (Dobson et al., 2012). Probiotics administered to tilapia (Oreochromis niloticus) increased lysozyme activity in host (Taoka et al., 2006).

Competitive exclusion of pathogenic bacteria
Probiotics often compete with pathogenic bacteria for space and nutrients that hinder their proliferation. Direct inhibition of pathogens by production of inhibitory compounds as discussed above is one way they competitively exclude pathogens. Other mechanisms include formation of biofilms, blocking adhesion sites and profuse probiotic growth. An oyster probiotic, P. inhibens S4 produces biofilms that inhibit the growth of pathogens V. coralliilyticus and V. anguillarum .
Exclusion of pathogenic bacteria by competition from probiotic bacteria was also shown in poultry. Native bacteria from adult chickens were used to protect chicks from infestation of Salmonella infantis (Rantala and Nurmi., 1973) as well as other enteropathogens (Schneitz, 2005). Porcine probiotics Lactobacilli and Bifidobacteria compete for attachment sites on epithelial cells and exclude pathogens in the intestine (Gross et al., 2008).

Enhancement of the Epithelial Barrier
Gut is in constant contact with a large number of bacteria and its integrity is often one of the most important barriers against invading pathogens. Increased expression of genes involved in tight junction signaling due to probiotic treatment reinforces this barrier (Anderson et al., 2010). Escherichia coli Nissle 1917 (EcN1917) has been shown to not only prevent disruption of the mucosal barrier by enteropathogenic E. coli, but also restore mucosal integrity (Anderson et al., 2010). Probiotics differentially modulate epithelial cell responses via activation or suppression of distinct signaling pathways in a strain-dependent manner (Llewellyn et al., 2017).

Immunity Effects on mucosal immunity
Mucus is made up of polymerized mucins that protect hosts from pathogens, enzymes, toxins, dehydration and abrasion (Hardy et al., 2013). Lactobacillus plantarum 299v and Lactobacillus rhamnosus GG have been shown to up-regulate production of MUC2 and MUC3 intestinal mucins that weakens the adherence of pathogenic Escherichia coli O157: H7 (Mack et al., 1999). Probiotics mediate modulation of mucin expression as a strategy for intestinal colonization of beneficial microbes to the host (Caballero-Franco et al., 2007). Mucus contains lysozymes, antimicrobial substances, antibodies and enzymes that have added benefits in controlling pathogenic invasion. Production of these substances can be modulated by presence of probiotics. Probiotic treatment led to increase in lysozyme production in Japanese flounder (Ye et al., 2011). Probiotic strains such as Lactobacillus GG, Bifidobacterium actis Bb-12 (Rautava et al., 2006) and Saccharomyces boulardii (Rodrigues et al., 2000) have been demonstrated to enhance IgA production and secretion.

Immunomodulation
Probiotic research shows mounting evidence of probiotic-host communication through pattern recognition receptors resulting in modulation on key signaling pathways such as NF-kB and MAPK to enhance or suppress activation and influence downstream pathways (Bermudez-Brito et al., 2012, Hardy et al., 2013, De et al., 2014. Probiotics and pathogens share PAMPs/MAMPs that can induce innate inflammatory pathways. Secondary and chronic exposure to probiotics induce suppressive /tolerogenic response that modulate NF-kB and MAPK pathways (Llewellyn et al., 2017). Effect in humans for some example probiotics is illustrated in Fig 3. L. casei CRL 431 interacts with epithelial cells through TLR2 and induces an increase in the number of CD-206 and TLR2 receptors in the cells involved in the innate immune response in humans (Vinderola et al., 2005). Lactobacillus stimulates TLR9 that induces cytoplasmic accumulation of ubiquitinated IkB and inhibition of NF-kB activation (Lee et al., 2006).
L. reuteri and L. casei engage with C-type lectin, prime dendritic cells and that lead to increased production of IL-10 (Smits et al., 2005). In contrast, L. reuteri strains DSM 17938 and ATCC PTA 4659 downregulates expression of TNF-a, TLR4 and NF-kB and upregulates IL-10 expression in rats (Bermudez-Brito et al., 2012). Along with the influence on innate immunity probiotics also have impacts on adaptive immunity (Hardy et al., 2013).
In addition, increase in phagocytic activity in probiotic fed Nile tilapia (Oreochromis niloticus) (Vieira et al., 2010) and increase in total hemocyte count and serum agglutination activity in probiotic fed and challenged marine shrimp (Sayed et al., 2011) are also documented. Probiotics have also been shown to confer protection against many cellular stresses, which include oxidative stress-mediated apoptosis (Llewellyn et al., 2017).
Thus, probiotic bacterial strains can be generalized to exert immune-activation,deviation or -regulation/suppression responses (Hardy et al., 2013). Selection of probiotic strains especially in combination along with prebiotics can have beneficial effects on hosts. However, it is crucial to gain full knowledge of their modulatory capabilities and formulate their use with careful consideration.

Goals of this study
There has been much progress in understanding immunity in mollusks especially in bivalves but we still lack knowledge of larval immunity in eastern oyster C. virginica.
There is also a dearth of understanding in the effect of bacteria on larval immunity.
The overall goal of this study was to understand the mechanism of action of probiotics B. pumilis RI06-95 and P. inhibens S4 against pathogen V. coralliilyticus RE22 and formulate them for use in the field.
Laboratory grown bacterial culture of B. pumilis RI06-95 was previously shown to protect C. virginica larvae from infection of V. coralliilyticus RE22 .
The first aim was to formulate the probiotic such that it can be effectively used in hatcheries and to test their efficacy. A series of formulations were prepared and tested in lab as well as in hatcheries to establish their efficacy.
The second specific aim was to understand the immunological response of C. virginica larvae to both probiotics B. pumilis RI06-95 and P. inhibens S4 in order to understand if immunomodulation is one of the mechanisms of action of these probiotics. Next generation RNA sequencing technology was used to obtain the transcriptomic response of C. virginica larvae to probiotics in a lab controlled and a hatchery environment to thoroughly investigate their effect on several larval genes at a time.
The third specific aim was to understand the immunological response of C. virginica larvae to pathogen V. coralliilyticus RE22 in order to understand its pathogenesis. To investigate this, larval transcriptomes generated post challenge with V. coralliilyticus RE22 were compared to control transcriptomes.    Zhang, L., Li, L., Guo, X., Litman, G.W., Dishaw, L.J., Zhang, G., 2015. Massive expansion and functional divergence of innate immune genes in a protostome. Sci. Rep. 5.

Introduction
The bivalve shellfish (oysters, clams, scallops, and mussels) industry is an important and rapidly expanding area of aquaculture production. The total landings for oysters, clams and scallops in United States alone valued at $859 million (NMFS 2016).
A primary requisite for the aquaculture of most bivalve shellfish species is an abundant, reliable, and inexpensive supply of seed/small juveniles (Helm et al. 2004 Given the absence of an adaptive immune system in bivalves allowing for the use of vaccines as disease prevention tools, the use of probiotics is one of the most promising management strategies for shellfish disease prevention and control (Elston 1998;Verschuere et al. 2000;Prado et al. 2010 We previously reported that marine Bacillus pumilus RI06-95, a producer of the antibiotic amicoumacin (Socha 2008), antagonized growth of the shellfish pathogen V.
coralliilyticus RE22 in vitro and protected eastern oyster Crassostrea virginica and bay scallop Argopecten irradians larvae against experimental challenge with V.
coralliilyticus RE22 (Karim et al. 2013b;Sohn et al. 2016a). It was also shown that daily treatment of larval rearing tanks in a hatchery with RI06-95 led to a decline in the levels of Vibrio spp. on tank surfaces and an increase in the survival of larval oysters when challenged with a pathogen (Sohn et al. 2016a). Bacillus spp., have shown promise as probiotic bacteria to improve host survival, growth, and development in aquaculture (Queiroz & Boyd 1998;Luis-Villaseñor et al. 2011;Martínez Cruz et al. 2012;Li et al. 2014 et al. 1995;Schisler et al. 2004;Salinas et al. 2006;Savini et al. 2010;Dagá et al. 2013). An appropriate formulation should offer several advantages in addition to host protection, including: the stabilization of microorganisms during distribution and storage; ease in handling and delivery of the product; protection of the microbes from adverse environmental factors; and safety to the aquaculture species. Therefore, the successful development of an appropriate probiotic formulation requires testing for efficacy, safety, and stability, especially in bivalve hatchery facilities.
Here, we evaluate three novel formulations of the candidate shellfish probiotic B. pumilus RI06-95. We determine storage and usage potential, and test each formulation along with fresh cultures of the same probiotic bacterium for safety and host protection in both laboratory and in semi-commercial scale hatchery experiments.
While all three formulations resulted in stable products with suitable shelf lives, only a spray-dried formula provided a high level of safety and efficacy desired for a commercially viable product. Our results demonstrate a safe, stable, and easy-to-use formulation for C. virginica larval aquaculture production.

Oyster larvae
Laboratory bacteria were maintained as stocks in 50% glycerol at -80 °C until use. They were cultured on yeast peptone with 3% NaCl (YP30) media (5 g L -1 of peptone, 1 g/L of yeast extract, 30 g/L of ocean salt (Red Sea Salt, Ohio, USA)) at 28 °C with shaking at 175 rpm as described in Karim et al. 2013a.

Granulated Product Formulation (RI-G)
Probiotic B. pumilus RI06-95 was incubated in 2.25% NaCl (YP22.5) broth (yeast extract 1 g/L, peptone 5 g/L, 22.5 g/L ocean salt, Instant Ocean) at 25 °C and 175 rpm. An initial culture was incubated for 2 d, then transferred to fresh YP22.5 and incubated for 4 d. The culture was partitioned into 50 mL sterile centrifuge tubes and centrifuged for 10 min at 2,350 × g. After centrifugation, cell pellets were transferred into a sterile petri dish (100 × 15 mm), and dishes were swirled with 2-3 mL culture media to ensure that the surfaces were completely covered in cells. The dishes were then covered with single ply, light duty paper (Kimwipes®) and placed in a convection oven to dry at 30 °C with constant airflow for 24-48 h, depending on initial volume. The dry cell mass was extruded through three particle size (40s, 80s, and 325s) USA Standard Sieve stainless steel screens (Cole Palmer, Illinois, USA), yielding products with average particle sizes of 43, 177, and 420 µm, respectively. The resulting granulated products were transferred into sterile glass vials and stored at 4 °C. For hatchery trials the granulated formulations were scaled up following the same formulation procedure as above except bacterial cultures were centrifuged at 9,300 × g for 10 min and the final cell pellet was dried at room temperature (22 ± 3 °C) for approximately 2 days.

Lyophilized Product Formulation (RI-L)
Probiotic B. pumilus RI06-95 was cultured from frozen stocks and then centrifuged as above. After discarding the liquid supernatant, 25 mL of Sugar Salt Solution (SSS) (2.5 g/L Instant Ocean, 200 mM sucrose, filtered deionized (DI) water (pre-filtered through a 0.2 µm filter)) was added to each tube, and the cell pellet was resuspended using a vortex. The re-suspended cells were frozen at -20 °C for 12 h, and then lyophilized for 48 h (Labconco FreeZone 4.5 lyophilizer, Kansas City, MO, USA).
The tubes were stored at 4 °C until use. 100 mM sucrose was used as a cryoprotectant during the lyophilization process. For hatchery trials, individual tubes with a single dose of formulation for a target dose of 5 × 10 4 CFU/mL for 100 L were prepared.

Spray-dried formulations (RI-SD)
Spray-dried formulations were prepared by Envera LLC (West Chester, PA) using a proprietary formula. Computer controlled fermentation vessels were used to grow the probiotic and pasteurized to make 100% spore-based product. After pasteurization, the probiotic was centrifuged and spray dried into a fine powder that can be easily hydrated with seawater. The final concentration of the probiotic in the formulation was 8.6 × 10 11 CFU/mg of powder. For the hatchery trial, tubes of the appropriate amount of formulation for a target dose of 5 × 10 4 CFU/mL in each 100 L tank were prepared. At the hatchery seawater was added to the tubes and mixed thoroughly. The mixed formulation was then added to the tanks daily during feeding.

Fresh culture controls (RI)
In order to determine the influence of the formulation process itself on the effectiveness of the probiotic in vivo, we tested simultaneous treatments of freshly cultured B. pumilus RI06-95 (cultures prepared as described in Sohn et al. 2016b) alongside formulated treatments in all lab and hatchery trials.

Viability and stability of formulated products
Viability and stability of each formulation was measured by counting colony forming units (CFU) on 2.5% yeast peptone agar plates using serial dilutions. Preformulation cell concentrations in CFU/mL were measured from culture aliquots directly before centrifugation. The RI-L product was re-suspended in 50 mL filtered sterile seawater (FSSW). The RI-G was suspended at 5 mg/mL in FSSW for 10 min and then vortexed for 1 min. The RI-SD was suspended using 0.1 g into 50 mL FSSW, followed by 10-fold serial dilutions. The percent cell viability in the formulations was calculated as follows: % Viability = [(sample formulation CFU/mL) / (pre-formulation CFU/mL)] × 100% RI-L was stored at 4 °C, while samples of RI-G were stored at either room temperature or 4 °C and RI-SD stored at room temperature. The stability of the formulated probiotics was measured immediately after formulation (t = 0) and 1, 2, 5, and 8 weeks after formulation, except RI-SD. Each assay was performed in triplicate.

Laboratory pathogen challenge experiments
Laboratory challenge assays were conducted following protocols outlined in (Karim et al. 2013a). Briefly, larval oysters were placed into six-well plates with 5 mL of filtered sterile sea water (FSSW, 28 psu). Probiotic treatments were added to the larvae at a concentration of 10 4 CFU/mL and incubated at room temperature with gentle shaking. After 24 h, the larvae were placed onto a 42 µm nylon mesh and washed gently using FSSW, then placed back into the original wells. Finally, V. coralliilyticus RE22 was added to each well, with the exception of the non-challenged controls, at a final concentration of 10 5 CFU/mL. Larval survival was quantified ~ 24 h after the pathogen was added using the neutral red technique . Survival was calculated by using the formula: Survival (%) = 100 × (number of live larvae/total number of larvae).
The relative percent survival (RPS) of probiotic pretreatment compared to the challenged control was calculated using the formula: RPS (%) = [1 -(% survival challenged control treatment / % survival challenged treatment)] × 100 as described in .

Hatchery trials
Hatchery experiments were conducted at Roger William University (RWU), Bristol, RI or the Aquaculture Breeding Center at the Virginia Institute of Marine Sciences (ABC), following standard operating procedures at each hatchery. For each trial, twelve 60 L (ABC) or 100 L (RWU) conical larval rearing tanks were used. We performed four independent trials at RWU between January 2014 and July 2016, and one trial at VIMS in June 2015 (Trial IV), testing each of the formulations at least once.
Each trial was initiated by adding 8 -10 larvae/mL (800,000 to 1,000,000 initial larvae) per tank 1-2 d post-fertilization to the conical tanks. Tanks were randomly assigned to treatments and probiotic formulations were added daily at the time of feeding mixed with the algal food. Larvae were kept in static conditions and tanks were drained-down every other day, cleaned, and re-stocked with fresh water. Treatments, number of tanks per treatment, and trial duration for each trial is shown in Table 1.

Larval survival and growth during hatchery trial
Data was collected at the time of selected drain-down events. Oyster larvae were passed through different sized mesh screens (35, 55, 75, and/

Laboratory pathogen challenge of probiotic-treated larvae from hatchery
An aliquot of larvae from each tank collected at selected drain-down events was transported to the laboratory at University of Rhode Island. Oysters (about 40 -50 larvae) were placed in six-well plates and then challenged with V. coralliilyticus RE22 at a final concentration of 10 5 CFU/mL following the methods described in the laboratory challenge section. Oyster larvae from Trial IV could not be challenged since very low number of oyster larvae were left in the probiotic treated groups at the hatchery.

Determination of levels of Vibrio spp in the hatchery
Total number of Vibrio spp. was evaluated using a plate count method on thiosulfate-citrate-bile salts-sucrose medium (TCBS, Difco) (Sohn et al. 2016a).
Samples were collected from water in the rearing tank (3 x 10 mL), tank surfaces (by swabbing), and oysters (about 1,000) when the tanks were drained. Swab samples (3 per tank) of tank surfaces (about 48 cm in length in total) were collected from each tank for all except Trial V. Each cotton swab was placed into a sterile Falcon tube containing 1 ml of FSSW and then mixed vigorously. Oyster larvae were rinsed with FSSW, homogenized using a sterile pestle, and suspended in FSSW. Ten-fold serial dilutions of each sample were prepared in triplicate, and then triplicate 10 µL of each dilution were plated on TCBS agar plates. After a ten-fold serial dilution, 10 µL samples of each of the dilutions were spotted evenly onto TCBS agar plates in triplicate for all except trial 5. The inoculated plates were incubated for 16 -20 h at 28 ˚C and the colony forming units (CFU) were calculated. Results are expressed as CFU/mL, where 1 mL corresponds to 1 mL of water in the tank, 1 mL of swab suspension, or 1 mL of water contacting about 1,000 larvae. Determination of Vibrio spp. levels could not be performed on larvae from trial IV due to very low numbers of surviving larvae.

Statistical Analysis
Larval oyster survival data were subjected to arcsine square root transformation prior to statistical analysis. The one-way analysis of variance (ANOVA) was used to determine significance between treatments within each time point. The two-way ANOVA was also used to determine significance between groups with time and treatment as factors. The Tukey's or Sidak's multiple comparison tests were used for post-hoc pairwise comparisons. A p-value < 0.05 was considered to be statistically significant.
Formulation cell viability data were analyzed by two-way ANOVA followed by Tukey's Test for each temperature and each time point. All statistical analyses were performed using Graphpad Prism, version 6.0 (Graphpad Software, Inc.). Differences were considered to be significant at values of p < 0.05.

Viability and stability of formulated products
The stability of the three formulated products was assessed after storage for 8 weeks (RI-G and RI-L) ( Figure 1) or 16 weeks (RI-SD) at ambient temperature. The three formulated products varied in their final CFU/ml following storage. RI-L and RI-G had similar pre-formulation concentrations of 1 × 10 8 CFU/mL and 1.27 × 10 8 CFU/mL, respectively. We observed a loss in viability immediately after the RI-G formulation process (data for RI-SD not available), and then again one week after storage at both 4 and 27 °C. However, we note strong stability after this initial loss. The spray dried formulation had a concentration ~200-250-fold higher at 2.65 × 10 10 CFU/mL 16 weeks post formulation.  Table 2). In only two instances was there a significantly higher protection by formulation against the pathogen challenge than the fresh culture (Table 2, L III and SD II, Fig. 2D and 2F respectively).

Effect of daily treatment with probiotics in the hatchery on larval growth and survival
Based on successful protection from pathogen challenges in laboratory trials, all three formulations were tested in a hatchery. Treatments included in each hatchery trial and the length of treatment is described in Table 1 survival. Thus, the SD-formulation is safe for use with oyster larvae in the hatchery.

Effect of daily treatment with probiotics in the hatchery on larval survival to bacterial challenge
Larvae from the hatchery experiments were tested for improved survival following challenge with V. coralliilyticus RE22. Since pathogens could not be introduced into the hatchery, larvae were collected and subjected to laboratory challenges as described in methods section. Larvae exposed to the granulated or lyophilized probiotics in the hatchery did not show significantly higher survival to a 24 h bacterial (V. coralliilyticus by the fresh culture of RI06-95 in this trial was 36 ± 6 % on day 12 (Table 3). On the other hand, trial V showed significantly improved survival both with RI-SD and fresh culture of RI06-95 as compared to controls (One-way ANOVA; p < 0.05; Figure 5 G).
Effect of daily treatment with probiotics in the hatchery on levels of total Vibrio spp.
In general, daily treatment of tanks with either formulation of B. pumilus RI06-95 did not lead to a significant decrease in the levels of total Vibrio spp. in water, tank surfaces, or oyster larvae as compared to control groups at each of the time points ( Figure 6 and Figure 7). High levels of variability were observed between tanks and trials within treatments. Interestingly, levels of Vibrios in the water were lower than 10 3 CFU/mL in Trial I ( Figure 6A) and none were detected on the tank surfaces during this trial ( Figure   6D). Trial I was performed in January, a month in which lower levels of Vibrios are present in coastal waters in the region (and therefore in water being pumped into the hatchery) (Duan & Su 2005, Parveen et al. 2008. Similarly, very low levels of Vibrios were found in Trial V in the water ( Fig 6G). Levels of Vibrios on tank surfaces and larvae were not measured during Trial V. Overall the results show that certain days probiotic treated tanks (formulated or fresh) show reduced level of Vibrios spp. as compared to control but there is no significant trend to specifically ascertain that effect.

Discussion
We outline three formulation protocols, a granulation process (G), a lyophilization process (L), and a commercial spray-dried process (SD). Variation in terms of success was achieved for each of the formulations, with the spray-dried formulation showing overall the best performance.
Granulation process: A traditional approach for formulating microorganisms is airconvective drying, which is a cost-effective process for the dehydration of RI-G showed protection in laboratory experiments and did not show any detrimental effect on the larvae in any of the laboratory assays. However, in the hatchery trial formulation treated larvae showed reduced survival as compared to control and freshly grown probiotic. It demonstrated protection from pathogen challenge in laboratory trials but was unsuccessful in doing so in the hatchery trial. Despite the favorable results from viability and storage of the granulation protocol, research on the granulated product was discontinued in this study due to a negative influence on survival of larval oysters in hatchery settings.
Lyophilization: The lyophilized formulation (L) did not significantly impact cell viability after the formulation process. Lyophilization has previously been investigated as a way of preserving and formulating Bacillus spp. as probiotic products (Henn et al. 2015). To ensure sufficient viability after freeze-drying, a disaccharide cryoprotectant such as sucrose or trehalose is typically added to provide structural support to cell membranes and proteins (Leslie et al. 1995). We successfully used sucrose at a concentration of 100 mM that provided stability and viability over time.
RI-L led to variable results in larval survival in hatchery trials. It failed to provide protection from pathogen challenge in the 2 out of 3 laboratory experiments and the hatchery trial. It produced no observable negative effect on water quality. Our results suggest that the addition of sucrose may be responsible for the negative impact on larval survival, as sucrose alone (without B. pumilus RI06-96) lowered larval survival in 2 of 4 trials where it was investigated. Because a wide range of bacterial taxa can readily use sucrose, we suggest that its addition to the formulated product may encourage antagonistic bacterial growth, and presents greater risks than advantages.
Spray drying: Of the three formulations tested, the commercially prepared spray dried formulation was found to maintain the highest concentration at room temperature over time while also showing no negative impact on larval oysters in the laboratory or in the hatchery trials. After 16 weeks at room temperature, the SD-product still contained >2.65 × 10 10 CFU/g. Previous research has shown that probiotic concentrations of Bacillus products at around 1 × 10 4 CFU/ml provide optimal performance (Karim et al. 2013a;Sohn et al. 2016a), meaning to reach a final target concentration of 1×10 4 CFU/ml in a 1,000 L commercial tank, only ~0.4 g of RI-695 would need to be added.
This would be extremely cost effective for use at a larger scale. Another added benefit of the formulation is its ease of use. The powder quickly suspends in seawater and is added to the tank very easily.
The spray-dried formulation was also shown to perform as well or better than freshly prepared B. pumilus RI06-95 in both laboratory experiments and hatchery trials.
In hatchery experiments, RI-SD showed no significant reduction in larval survival, water quality or larval growth. In fact, it increased survival compared to freshly prepared culture in the hatchery trial by day 12. RI-SD also performed well in pathogen challenge experiments, increasing survival of larvae after the challenge at the same rate or greater as compared to freshly prepared culture.
As seen in previous hatchery experiments , high levels of variability were seen between tanks and trials within a treatment. The variation in results within and/or between experiments in this study could be due to several factors: (1) a different quality and health status of larvae from each trial; (2) the impact of various environmental and biological factors such as salinity, pH, temperature/season at the hatchery; (3) variability in the characteristics of different rearing systems, such as tank, source or treatment of water, and location of hatchery (Balcazar et al., 2006;Gatesoupe, 1999;Martínez Cruz et al., 2012;Utting and Millican, 1997); and 4) the effect of variability in the composition of microbial communities and how these communities may interact with the probiotic. Due to above factors the variability is more pronounced in hatchery trials than controlled laboratory experiments. Although variability is seen within and/or between experiments in this study for RI-G and RI-L, it is highly minimized in the trials using RI-SD. More importantly there is consistency in the goal of achieving protection from pathogen challenge with use of RI-SD.
The use of probiotics as a disease control mechanism has particular and critical relevance to shellfish hatcheries, where disease losses are high, vaccination is not possible and use of antibiotics is not recommended. Our results demonstrate successful formulation of the candidate probiotic B. pumilus RI06-95 for its use in shellfish hatcheries using the spray drying method. It also demonstrates the challenge in formulating the probiotic and the need of thorough testing in both laboratory and hatchery setting to confirm the effect of formulation. The laboratory and hatchery trials confirm that the RI-SD formulation is stable over a long term, remains viable and shows comparable performance to freshly grown cultures of the probiotic. It is suitable for storage, transportation and can be easily applied in a hatchery by mixing with sea water.
Although the addition of RI-SD did not show reduction in Vibrio spp. in general, this might not be a strategy used by the probiotic as its mechanism of action. Probiotics are known to modulate the immune system of the host (Hardy et al., 2013, Mortha et al., 2014, Sanchez et al., 2015. This could be one the strategies used by B. pumilus RI06-95 to provide protection in the event of vibriosis. Future research in mechanism of action of the probiotic would help in optimization of the use formulation in terms of dosage timing and frequency. Figure 1. Impact of formulation processing (granulation or lyophilization) and temperature storage on the stability of Bacillus pumilus RI06-95. Cell count in the reconstituted formulation after storage for up to 8 weeks was determined using a plating method. Data expressed as mean ± SEM of CFU/mg of formulation.

CFU/mg of formulation
probiotic, control for lyophilized formulation); RI-G = granulated formulation; RI-L = lyophilized formulations (in 100 mM sucrose); RI-SD = spray-dried formulation; RI = RI06-95 freshly cultured in lab.      showed evidence of suppression of key immune signaling pathways but possibly activated antiviral pathways. The larval response to RE22 lacked production of protease inhibitors, hypothesized to be involved in providing protection against the proteases that are a key virulence factor of RE22. In addition, transcriptomic data suggests modulation of mucus and cytoskeletal components. The transcriptomic response was also characterized by differential expression of metabolic genes, suggesting high metabolic demand and oxidative stress contributing to larval mortality. This study fills a major gap in our knowledge on the immune responses in larval stages of this economically and ecologically important species. This information could aid in developing solutions to control disease and design better management practices for hatcheries.

Introduction
Vibrio spp. are common pathogens causing disease in a wide variety of aquatic species, including several species of mollusks. Strains of V. coralliilyticus also cause disease in corals, leading to bleaching (Ben-Haim et al., 2003, Wilson et al., 2013. V. coralliilyticus RE22, previously known as V. tubiashii RE22, causes vibriosis in bivalve larvae (Richards et al., 2015). The disease resulted in heavy mortalities that severely affected oyster seed production of shellfish hatcheries (Elston et al., 2008).
Infection by vibrios in bivalve larvae is dramatically rapid in progression and characterized with signs of bacillary necrosis, reduced feeding, and swarming of bacteria around the moribund larvae (Tubiash et al., 1965). An investigation of the colonization and infection process in Manila clam (Ruditapes philippinarum) larvae using a GFP-tagged Vibrio sp. showed pathogen entry through ingestion, with infection quickly spreading to other organs and followed by colonization and proliferation in the entire body (Dubert et al., 2016). The genome of V. coralliilyticus RE22 shows that it encodes several extracellular metalloproteases, serine proteases, hemolysins and type secretion systems as virulence factors (Hasegawa et al., 2008, Hasegawa et al., 2009. Several studies have characterized changes in gene expression patterns in larval stages of bivalves during development including Pinctada fucata (Li et al., 2016), C. angulata (Qin et al., 2012) and in response to vibrio infection in Crassostrea gigas (Hasegawa et al., 2008) and C. virginica (Genard et al., 2012). This study aims to enhance knowledge on bivalve-vibrio interactions by analyzing the transcriptomic response of larval eastern oysters, an economically and ecologically important species, to infection with V.
coralliilyticus RE22, a bacterial pathogen capable of causing rapid and high levels of mortality in bivalve hatcheries. The goals of this study are to (1) characterize the response of C. virginica larvae to experimental challenge with V. coralliilyticus RE22; and (2) provide hypotheses on possible strategies used by V. coralliilyticus RE22 to overcome larval immune defenses. This information will aid in developing solutions to control disease and design better management practices for hatcheries.

Vibrio coralliilyticus RE22 culture:
The pathogen (supplied by H. Hasegawa, Department of Biomedical Sciences, Oregon State University) was maintained and stored in 50 % glycerol stocks at -80°C until use.
Inocula from freezer stocks were plated on yeast peptone with 3%NaCl (YP30; 5 g L -1 of peptone, 1 g L-1 of yeast extract, 30 g L-1 of ocean salt, Instant Ocean) agar plates for 2 d, then transferred to 5 mL of YP3 broth incubated at 25°C on a shaker (134 rpm) for 1 d. Cultures were washed using Artificial Filtered Sterile Seawater (AFSW, 28-30 psu salinity) twice by centrifugation at 23,000 rpm for 10 min. The OD at 550 nm was measured and the stock was diluted such as to obtain a sub lethal concentration of 5 × 10 4 CFU mL-1 for transcriptome analysis and a lethal concentration  of 5 × 10 5 CFU mL-1 for disease progression analysis.

Oyster larvae:
C. virginica larvae were obtained from shellfish hatcheries on the east coast of United States. Larvae 6-10 days old were collected at the hatchery and shipped overnight to the lab at the University of Rhode Island on a wet filter. Upon arrival to the laboratory, larvae were washed with AFSW on top of a 40 µm nylon mesh and placed in stock containers containing AFSW. Larvae were acclimatized to the laboratory environment (room temperature) for 24 h prior to the experiments.

Effect of V. coralliilyticus RE22 on mortality of C. virginica larvae
In order to understand the rate of progression of disease, C. virginica larvae were experimentally challenged with 5 x 10 5 CFU mL -1 V. coralliilyticus RE22 . Larval density (larvae/mL) of the stock was determined using a Nikon E200 microscope. Larvae (~100) were distributed in wells of a 6-well plate with 5 mL AFSW and maintained at 22 -23 °C with gentle rocking. Two treatments (control and challenge) were each conducted in triplicate. Larval mortality was recorded at 6, 9, 14, 18 and 20h post addition of V. coralliilyticus RE22 by evaluation of active swimming and/or gut and cilia movement using a Nikon E200 microscope.

Experimental set up
For biological replicates, the complete set up as explained below was performed using larvae from three different hatcheries (n = 3 experiments, operationally designated as K, M, and V). Larvae from the stocks were distributed into tissue culture flasks (~10,000 per flask) in 500 mL AFSW and kept on a shaker with gentle shaking at ~50 rpm at room temperature. Larvae were acclimatized to the experimental set up for an additional 24h prior to challenge. Each treatment (control and challenge) was conducted in duplicate to serve as technical replicates. Larvae were fed with 1 mL of instant algae Shellfish Diet 1800 TM (20,000 cells/mL; Reed Mariculture Inc., San Jose, CA. USA) immediately prior to treatment in order to promote pathogen ingestion. Challenge with V. coralliilyticus RE22 was performed with a sub lethal concentration of 5 × 10 4 CFU mL -1 for 6h.

Larval Collection post treatments
Control larvae were collected at 0h and RE22 treatments were collected 6h post challenge. Larvae were aspirated gently from the flasks using a 100mL serological pipette, and filtered through a 40 µm sterile filter for collection. Since dead larvae settle to the bottom, the last 25 mL of each flask was not collected to avoid bias in transcriptomic response. Larvae were washed with 2mL of AFSW on a 40µm filter, followed by a wash using 2mL of RNAlater™, aspirated from the filter using a pipette, placed in labeled 2 ml RNase free microfuge tubes, and held at 4°C for 24h in RNAlater™ followed by storage at -20°C.

RNA extraction, cDNA prep and sequencing
Tri-reagentä (Sigma-Aldrich) was used for extracting total RNA from all the samples following manufacturer's instructions (TRI Reagentä Protocol, Sigma-Aldrich). RNA extracts were DNase treated using the DNA-freeä DNA removal kit from Ambion and purity and concentration of RNA was assessed using a Nanodrop 8000 spectrophotometer (Thermo Scientific

Assembly, annotation and analysis
Raw reads obtained from sequencing were filtered, trimmed and adapters were removed using bbduk program in BBTools suite from Joint Genome Institute and viewed in FASTQC (Andrews, 2010). Processed reads were aligned to C. virginica reference genome (version 3.0) via HISAT2 2.1.0 (Kim et al., 2015) and assembly was performed using Stringtie (Pertea et al., 2016) using default parameters. To compare the depth of sequencing across all samples preseq package was used (Daley and Smith., 2013).
Differential gene expression analysis was performed by comparing transcript counts between RE22 6 h treatment (replicates K, M, V) vs control 0 h (replicates K, M, V) using DESeq2 (Love et al., 2014). Transcripts with Benjamini-Hochberg adjusted p value £ 0.05 and log fold change of ≥ 2 or ≤ -2 were considered significantly differentially expressed. This analysis design only allowed for the most conservative estimates and only showed differentially expressed genes representing all the biological replicates. Annotation for differentially expressed genes (DEGs) was performed by mapping to NCBI protein non-redundant (NR) database using BLASTx (Altschul et al., 1997) with an e-value cutoff of 1e -3 and hit number threshold of 20. Mapping DEGs to GO terms was conducted using BLAST2GO v4.1.9 (Conesa et al., 2005) and functional enrichment was done using topGO (Alexa et al., 2006) with default parameters. ReviGO (Supek et al., 2011) was used to plot and visualize results obtained from topGO with default parameters (allowed similarity was set to medium). Significantly enriched GO terms were obtained by using Fishers exact test (p £ 0.01). Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway annotations were also obtained using the KEGG Automatic Annotation Server (KAAS).

Effect of V. coralliilyticus RE22 on mortality of C. virginica larvae
Mortality in larval oysters exposed to 5´10 5 CFUmL of V. coralliilyticus RE22 was initially seen at 14h after challenge, increasing exponentially after that (Figure 1a). The larvae appeared normal at 6h, but 9h after challenge many showed reduced motility and feeding (Figure1b).

Transcriptome alignment
Depth of sequencing for all the transcriptomes ranged from 16,617,375 -39,681,499 paired end reads. Sequencing saturation curves for all transcriptomes were close to full saturation, indicating that all but the rarest transcripts would be represented in the transcriptome ( Figure 2). The alignment rate to the Crassostrea virginica reference genome using HISAT2 ranged from 85 -89 % (Table 1).

Differential expression Analysis
Comparison of transcriptomes obtained from RE22 treated (6h) larvae to control (0h) larvae using DESeq yielded 1,534 differentially expressed transcripts (p ≤ 0.05, upregulation: log fold change ≥ 2, downregulation: log fold change ≤ -2). Refer to supplementary data tables in appendix for descriptions and log fold change values.

GO and KEGG annotation
A Gene Ontology (GO) term enrichment analysis was performed on all the differentially expressed transcripts in response to RE22 challenge. There were 22 biological processes significantly enriched (p<0.05) that mainly belonged to metabolism and signaling, but none related to immunity (   (Table 3).

Immune related genes
Described below are some of the important immune-related genes showing differential expression in RE22 exposed larvae (6h) as compared to control (0h) ( In terms of immune effectors, some mucin transcripts were differentially expressed in response to RE22, showing a mixed response (both up and downregulation).
In addition, cytoskeleton related transcripts including cytoplasmic actin and septin-11like were downregulated, but dynamin-1-like transcripts showed high levels of upregulation.

Cell death
Transcripts corresponding to autophagy related gene ATG9A were highly upregulated in response to RE22. Several transcripts that belong to the apoptosis pathway were differentially expressed in response to RE22 including transcripts identified as death domain-containing protein CRADD-like, caspases (1, 2, 6, 7-like) and IAP3 were upregulated while caspase 3 and IAP2 were downregulated (Table 4).

Metabolism and oxidative stress
Transcripts involved in metabolism that were differentially expressed included Cyt p450 and Cyt c subunits I and III. Heat shock proteins HSP12A and HSP12B were highly upregulated, while a few limited antioxidant enzymes were upregulated in response to RE22 (Table 4).

Discussion
Both differential expression and functional enrichment analyses of oyster larvae 6h after

Differentially expressed immune genes in response to RE22
Highlights of the immunological response of C. virginica larvae to V. coralliilyticus RE22 at 6h of exposure include pathogen detection via activated pathogen recognition receptors. However, along with an increased expression of immune receptors, an overall suppression of key immune signaling pathways and lack of specific immune effectors against RE22 was seen, suggesting that the pathogen is able to neutralize the immune response of the larval host.
Pattern recognition receptors (PRRs) are extremely important to innate immune system that recognize conserved pathogen-associated molecular patterns (PAMPs) and trigger signaling pathways that produce a variety of antimicrobials (Akira et al., 2006).

Conflicting immune gene responses:
Along

Antiviral immune gene responses:
Although, differentially expressed transcripts in response to RE22 indicate majority of the key immune signaling pathways to be suppressed, antiviral pathways seem to remain active. STING is a key regulator for sensing intracellular single-or double-stranded nucleic acids. STING via the cGAS-STING pathway complex with TAK1 and trigger expression of interferon genes. cGAS is activated whenever foreign DNA (both bacterial and viral nucleic acids) is detected in the cytoplasm (He et al., 2015, Gerdol, 2017. These results suggest the possibility of an intracellular invasion by RE22 that could lead to activation of STING or effectors of type secretion systems of RE22 (T6SS or T1SS) inadvertently leading to activation of these pathways. A special STING homolog LvSTING was activated in shrimp in response to V. parahaemolyticus infections that participates in antimicrobial peptide production .
Similarly, activation of JAK-STAT pathway has been reviewed in bivalves as an antiviral response (Green et al., 2015) but microbial activation of this pathway has been shown in Chinese mitten crab Eriocheir sinensis (Li et al., 2013).

Effectors
Extracellular metalloproteases in V. coralliilyticus RE22 are shown to be important in its pathogenicity to C. gigas larvae (Hasegawa et al., 2008). Therefore, the observation of lack of serine protease inhibitors in challenged larvae, as well as the lack of upregulation of other types of protease inhibitors, was unexpected. It is possible that protease inhibitors are not differentially expressed at the time point tested (6 h Mucus is the first line of defense in oysters besides the closed oyster shell. Mucus was one of the few immune effectors shown to be upregulated in larval oysters exposed to V. coralliilyticus RE22. Some pathogenic Vibrio spp. require binding to mucin in the gut epithelium as a part of their pathogenesis (Bhowmick et al., 2008, Jang et al., 2016, so it is possible that modulation of host mucus production or composition may allow RE22 to bind better and breach host defenses in larval oysters.

Cytoskeletal reorganization
Downregulation of septin-11 associated with the cytoskeleton in response to RE22 suggests possible disruption of cytoskeleton by RE22, but the functional implications of this downregulation is not clear. Both actin and septin 8B were shown to be upregulated by challenge with V. splendidus LGP32 in C. gigas (Duperthuy et al., 2011) and soft-shell clams, Mya arenaria  for hemocyte invasion.
Cytoskeletal disruption using upregulation of ß-actin due to V. tapetis challenge in Ruditapes philippinarum has also been demonstrated (Brulle et al., 2012). We need to know more about nature of RE22 pathogenesis in cytoskeletal modulation to fully understand this.

Cell death
It is difficult to interpret whether apoptosis is inhibited or enhanced in response to RE22 treatment due to modulation of both pro (caspases) and anti-apoptotic (apoptosis inhibitor) genes. This was also the case in surviving C. gigas on exposure of different strains of virulent Vibrio spp. . IAPs were modulated in both susceptible and resistant C. virginica families in response to A. crassostreae . The mechanisms underlying pathogen-induced modulation of apoptosis in mollusks are not well understood.

Metabolism and oxidative stress
Differential expression of heat shock proteins and cytochrome oxidases during RE22 challenge suggests that larvae are experiencing stress and high metabolic demand due to the inability to rapidly clear RE22 infection. Higher stress levels and lower metabolic rates have been seen in late responses (24-48h) of V. coralliilyticus LPI 06/210 in C.
gigas (Gernard et al., 2013). Moreover, no increase in expression of antioxidant enzymes, necessary to deal with oxidative stress from activated metabolism, was seen in our study. These results suggest that oyster larvae, which already possess a high metabolic demand to sustain the processes of rapid growth and development, may not be able to cope with the additional metabolic demand associated with infection.
Moreover, reduced feeding in infected moribund larvae may not allow replenishment of energy to mount an expensive immune response (Gernard et al., 2011). It has been shown for several V. tubiashii strains affecting bivalves that the pathogen enters the host through ingestion, proliferates in the gut, then spreads to other organs, including the cilia that are involved in swimming and capturing particles (Tubiash et al., 1965).

Conclusion:
The observed absence of induced expression of protease inhibitors, antimicrobial peptides or other immune effectors able to block RE22 virulence factors, along with other indications of a suppressed immune system, suggest that larvae are left highly susceptible to disease and then succumb to infection. Additionally, differential gene expression analysis indicative of a high metabolic demand and oxidative stress are consistent with the rapid mortality observed during RE22 infection in oyster larvae.

Abstract
The eastern oyster Crassostrea virginica is an ecologically and economically important species. Bacterial pathogens like vibrios cause heavy mortalities in oyster larvae in hatcheries. Probiotics are an inexpensive, practical, and natural method of disease control. Pretreatment of larval oysters with probiotics Bacillus pumilus RI06-95 and Phaeobacter inhibens S4 significantly decreases mortality caused by experimental challenge with the pathogen Vibrio coralliilyticus RE22. The aim of this study was to understand the oyster larval immune response to probiotics RI06-95 and S4 and the role it may play in protecting larvae from pathogen challenge. C. virginica larvae were exposed to each probiotic in two settings: controlled 6 and 24 hours laboratory exposures and 5 to 16 days exposure in a hatchery. Transcriptomes were sequenced using high throughput RNA sequencing and aligned to the C. virginica reference genome. Differential expression analysis compared probiotic treated transcriptomes to unexposed controls. Key features of the host immune response were shared despite the length of probiotic exposure, type of probiotic exposure and the type of environment in which exposures were conducted. Transcriptome analysis showed increased expression of genes for receptors involved in environmental sensing and detection of pathogens, immune signaling pathways, and immune effectors including serine protease inhibitor, mucins and perforin-2. In addition, patterns of differential gene expression suggest that inhibition of apoptosis, enhanced autophagy, and cytoskeletal reorganization may play a supplemental role in bacterial clearance. Thus, results from this study suggest that larval oysters show a robust and effective immune response to probiotic exposure, contributing to clearance of the probiotic within 24 hours after exposure. Activation of antibacterial immune effectors by probiotics, when provided 6 -24 hours prior to bacterial challenge, may play an important role in protecting larvae from mortality by V. coralliilyticus RE22. However, for continued effective protection, probiotics should be applied repeatedly and for at least 6 hours prior to RE22 challenge. This is the first time that immune responses of larval stages of C. virginica to bacteria are studied using a larval transcriptome. This research provides important new insights into host-microbe interactions in larval oysters that could be applied in the design of improved strategies for use of probiotic organisms for disease control in hatcheries.

Introduction
The eastern oyster Crassostrea virginica is an economically and ecologically important species (Newell 2004, NMFS 2014. Rearing of oyster larvae is a critical step to ensure a healthy and sufficient supply of seed for aquaculture industry. Bacterial diseases commonly described in larval stages are associated with high mortalities in hatcheries (Lauckner et al. 1983, Sinderman et al. 1990). Vibriosis is one such disease that leads to mortality in oyster larvae and juveniles (Tubiash et al, 1965). Bacteria of the genus Vibrio are ubiquitous within marine environments and detected in tissues of many marine organisms including abalones, bivalves, corals, fish, shrimp, sponges, squid, and zooplankton ( Thompson et al. 2004). Vibrio can cause larval mass mortalities in hatcheries in a short period of time leaving few options for treatment (Helm and Lovatelli 2006). In order to eliminate Vibrios and sanitize the facility, hatcheries need to shut down for several days after a disease outbreak before production is resumed (Helm and Lovatelli 2006). In particular, V. coralliilyticus RE22 (previously V. tubiashii RE22) has caused high larval and juvenile mortality in hatcheries (Elston et al. 2008). Vibrios are known to produce potent exotoxins that affects larval motility in oysters. Incapacitated ciliary movement affects feeding, leading to death due to starvation (DiSalvo et al., 1978, Brown and Roland, 1984, Kennedy, 1996. The extracellular metalloprotease secreted by V. coralliilyticus is toxic and induces mortality in oyster larvae (Hasegawa et al., 2008).
Practices to reduce mortality due to bacterial disease include treatment with antibiotics and disinfection of seawater. Water treatment, however, is expensive and could be toxic to the larvae if not properly done, while antibiotic treatment can lead to bacterial resistance. Treatment with antibiotics raises environmental and human health concerns as well (Prado et al. 2009, Akinbowale et al., 2016, Ho et al., 2000. Therefore, alternative methods need to be developed to manage good larval rearing environment and to control bacterial diseases in bivalve shellfish hatcheries.
Probiotics are defined as a live microbial food supplement that, when administered in a sufficient amount, confers a health benefit on the host (Food and Agricultural Organization of the United States 2006). Probiotics are known to benefit the host by a variety of means, including production of antimicrobials, improving water quality, enhancing the immune responses of host, and competing for space with pathogenic bacteria (Verschuere et al. 2000). There is growing evidence that probiotics show immunomodulatory effects in fish and shellfish (De et al., 2014, Newaj-Fyzul et al., 2015. The benefits of probiotics have already been shown in Pacific oysters, Crassostrea gigas (Douillet and Langdon 1994) and the eastern oyster C. virginica ).
Pretreatment of larval and juvenile C. virginica with probiotic organisms Phaeobacter inhibens S4 (isolated from the inner shell of oysters) (referred to as S4) and Bacillus pumilus RI06-95 (isolated from a marine sponge from the Narrow River in Rhode Island) (referred to as RI) before exposure to the bacterial pathogens Alliroseovarius crassostreae and Vibrio coralliilyticus RE22 (referred to as RE22) improves oyster survival rate . Additionally, probiotics are not harmful to oysters in absence of pathogens .
S4 is a Gram-negative organism and production of the antibiotic tropodithietic acid (TDA) and biofilm formation are two mechanisms utilized by S4 for protecting oysters from infection. Mutants of S4 unable to produce TDA and with decreased ability to produce biofilms, however, still provide some level of protection  suggesting that other mechanisms are also potentially involved. RI is a Gram-positive organism and produces the antibiotic amicoumacin, but this antibiotic does not inhibit the growth of RE22 in an in vitro assay, indicating that other mechanisms of action are also likely involved in RI's protection of larvae against bacterial challenge . Probiotics are known to act as immunomodulators (Hardy et al., 2013, Mortha et al., 2014, Sanchez et al., 2015.  Vidal-Dupiol., et al., 2014). Similarly, immune response of soft-shell clams, Mya arenaria, to V. splendidus strain LGP32 showed an overall downregulation of immune genes such as ficolin, killer cell lectin-like receptor, natural resistance-associated macrophage protein 1 (Nramp-1), and mitogen-activated protein kinases (MAPK) . Our hypothesis is that pre-treatment of oyster larvae with probiotics may cause an activated immune state in larvae that would serve to counteract the immunosuppressive effects of RE22.
Not much is known about the impact of friendly or beneficial bacteria on the immune system of oysters. The goal of this study is to determine the immunological response of C. virginica larvae to exposure to two probiotic bacterial species that differ in Gram character, in order to understand the potential role of immunomodulation as a potential mechanism of action of the probiotics in providing protection against V. coralliilyticus RE22.

Probiotic Bacterial strains:
Probiotic isolates S4 and RI were maintained and stored in 50 % glycerol stocks at -80°C until use. Bacteria were cultured by plating out freezer stocks on yeast peptone with 3% NaCl (YP30) agar plates for 1 d then transferred to 5 mL of YP30 broth (5 g L -1 of peptone, 1 g L -1 of yeast extract, 30 g L -1 of ocean salt, Instant Ocean) incubated at 28°C on a shaker (134 rpm) for 2 d. Cultures were washed using Artificial Filtered Sterile Seawater (AFSW, 28 -30 psu salinity) twice by centrifugation at 23,000 g for 10 min. The OD at 550 nm was measured and the stock was diluted to obtain a concentration of 5 × 10 4 colony forming units (CFU) mL -1 as previously described .

Oyster larvae:
C. virginica larvae were obtained from three shellfish hatcheries on the east coast of United States including Oyster Seed Holdings, VA, Virginia Institute of Marine Science, VA and Aeros Cultured Oyster Company, NY that served as three biological replicates. Larvae 6-10 days old were collected at the hatchery and shipped to the laboratory at the University of Rhode Island on a wet filter overnight. Upon arrival to the laboratory, larvae were washed with AFSW on top of a 40 µm pore size nylon filter to prepare a stock. The stock of larvae from each hatchery was used for probiotic exposures as described below. The same stock was also used for characterizing immune response to pathogen V. coralliilyticus RE22 (Modak et al., in prep, Chapter 3 of this dissertation).

Effect of length of probiotic pretreatment on protection against bacterial challenge
Previous research on the effect of probiotics on protection against challenge with the bacterial pathogen V. coralliilyticus RE22 was performed using a 24 h pre-incubation period with the probiotics prior to bacterial challenge . In order to determine if a shorter pre-incubation period with probiotics would confer protection against bacterial challenge, ~100 larvae were placed in each well of a 6 well plate in 5 mL of AFSW and incubated with 10 4 CFU mL -1 of probiotics S4 or RI06-95 for 6 or 24h prior to bacterial challenge with 10 5 CFU mL -1 of RE22. Larval survival was determined 24 h after challenge using previously described methods .
Survival rate was calculated as follows: Survival rate (%) = 100 x (number of live larvae/total number of larvae). One-way analysis of variance (ANOVA) was used to determine significance between treatments and Tukey's multiple comparison tests were used for post-hoc pairwise comparisons (p < 0.05) .

Effect of short-term exposure to probiotics on larval gene expression 2.4.1 Experimental set up for laboratory-scale experiments:
For biological replicates, three independent experiments were performed using larvae from three different hatcheries. Larval density (larvae mL -1 ) of the stock was determined using the Nikon E200 microscope. Two parallel exposures were performed with each set of larvae: (i) a large-scale incubation for collection of larvae for transcriptome analysis, and (ii) a small-scale experiment in 6 well plates for evaluation of the effect of probiotic exposure on protection against bacterial challenge. Probiotics were applied at a concentration of 10 4 CFU mL -1 . Larvae were fed with instant algae Shellfish Diet 1800 TM (20,000 cells/mL; Reed Mariculture Inc., San Jose, CA. USA) just prior to treatment in order to promote probiotic ingestion.
(ii) Set up for verification of protection by probiotics: Oyster larvae (~100) were placed in each well of a 6 well plate in 5 mL of AFSW and incubated with 10 4 CFU mL -1 of probiotics S4 or RI for 6 or 24 h prior to bacterial challenge with 10 5 CFU mL -1 of RE22, as described in section 2.3 above.

Larval Collection post treatments:
After incubation with probiotics, larvae from the flask set up for the transcriptome experiment were aspirated gently using a 100 mL serological pipette and filtered through a 40 µm sterile filter for collection. Since the dead larvae settle to the bottom, the last 25 mL of each flask was not collected to avoid bias in transcriptomic response.
Larvae were washed with 2 mL of AFSW followed by 2 mL of RNAlater™. Larvae retained on the filter were aspirated with a pipette using 1.5 mL of RNAlater, placed in labeled 2 mL RNase free microfuge tubes, and held at 4°C for 24 h in RNAlater followed by storage at -20°C.

RNA extraction, cDNA prep and sequencing:
Tri-reagent™ (Sigma-Aldrich) was used for extracting total RNA from all the samples following manufacturer's instructions (TRI Reagent™ Protocol, Sigma-Aldrich). RNA extracts were DNase treated using the DNA-free™ DNA removal kit from Ambion and purity and concentration of RNA was checked using a Nanodrop 8000 spectrophotometer (Thermo Scientific). Technical replicates were pooled at equimolar concentration. The quality and quantity of the pools were assessed using Agilent 2100 Bioanalyzer and High Sensitivity D1000 ScreenTape®. RNA samples were selectively enriched for poly-A containing mRNA and cDNA libraries were prepared using the PrepX RNAseq library Prep Kit (Takara Bio USA, inc). Samples were sequenced on Illumina HiSeq platform with 2×125 reads and sequencing coverage of 20-30M per sample at the Harvard University, FAS Center for Systems Biology, MA.

Effect of exposure to probiotics in the hatchery on larval gene expression 2.5.1 Experimental set up of hatchery experiments:
Transcriptomes obtained from treatment of larvae with B. pumilus RI06-95 will be referred to as HT_RI. Adult eastern oysters were spawned at the Blount Shellfish Hatchery, Roger Williams University, RI. Each trial was initiated by adding 8-10 larvae mL -1 (800,000 to 1,000,000 initial larvae) per tank 1 day post-fertilization. Larval oysters were distributed into 100 L conical tanks filled with filtered and UV treated seawater (20 -24 C and 28 -30 psu salinity) 1 day after fertilization and fed live microalgae daily from a microalgae production greenhouse. Water from Narragansett Bay, RI was filtered and UV treated and used for the larval tanks. Treatments included control and probiotic RI treated at a concentration of 10 4 CFU mL -1 . Each treatment was conducted in triplicate. Probiotics were added daily at the time of feeding.

Larval Collection post treatments:
Larvae for transcriptomes were collected at three time points: 5, 12 and 16 days post fertilization from probiotic-treated and control tanks. Larvae had been treated with probiotics daily starting 1 day after fertilization, as described in .
Tanks were drained on a filter with suitable pore size (75 -150 µm depending on the age of the larvae) at the time of collection. Using a serological pipette, larvae were aspirated gently and collected in RNase free microfuge tubes with RNAlater™ and stored at -80°C.

Verification of protection by probiotics:
A subsample of larvae was collected from each treatment and control tanks on day 8 post-fertilization to determine the effect of exposure to the probiotics in the hatchery on protection against bacterial challenge.
Levels of protection were determined using the methods described in 2.3. above, with the following modifications: larvae from each tank were placed in triplicate wells in 6well plates with ~100 larvae per well V. coralliilyticus RE22 at 10 5 CFU mL -1 dose.

RNA extraction, cDNA prep and sequencing:
Larvae were processed for RNA extraction as described in 2.4.4 above. cDNA libraries were generated using random hexamer priming that were sequenced on Illumina HiSeq platform with 2×150 reads and sequencing coverage of 50-70M per sample at the McDonnell Genomics Institute, Washington University School of Medicine, MO.

Assembly, annotation and analysis
Raw reads obtained from sequencing were filtered, trimmed and adapters were removed using bbduk program in BBTools suite from Joint Genome Institute and viewed in FASTQC (Andrews, 2010). Processed reads were aligned to C. virginica reference genome (version 3.0) via HISAT2 2.1.0 (Kim et al., 2015) and assembly was performed using Stringtie (Pertea et al., 2016) with default parameters. To compare the depth of sequencing across all samples preseq package was used (Daley and Smith., 2013).
Differential gene expression analysis between probiotic (RI or S4) treatment at each time point (6 or 24 h) and control (time 0, common to all treatments) was performed using DESeq2 (Love et al., 2014) and transcripts with Benjamini-Hochberg adjusted pvalue £ 0.05 and log fold change ≥ 2 or ≤ -2 were considered significantly differentially expressed. For hatchery transcriptomes, each of the days (5, 12 and 16) were considered as biological replicates and an overall comparison of treatment vs control was conducted. Transcript counts for each replicate were used to determine which DEGs are present in each replicate individually. This analysis design only allowed for the most conservative estimates and only showed differentially expressed genes representing all the biological replicates. Annotation for differentially expressed genes (DEGs) was performed by mapping to NCBI protein non-redundant (NR) database using BLASTx (Altschul et al., 1997) with an e-value cutoff of 1e -3 and hit number threshold of 20. Mapping DEGs to GO terms was conducted using BLAST2GO v4.1.9 (Conesa et al., 2005) and functional enrichment was done using topGO (Alexa et al., 2006) with default parameters. ReviGO (Supek et al., 2011) was used to plot and visualize results obtained from topGO with default parameters (allowed similarity adjusted to medium). Significantly enriched GO terms were obtained by using Fishers exact test (p £ 0.01). Kyoto Encyclopedia of Genes and Genomes (KEGG) pathway annotations were also obtained using the KEGG Automatic Annotation Server (KAAS).

Effect of length of probiotic pretreatment on protection against bacterial challenge
A short duration of S4 or RI pretreatment (6 h) showed variable levels of protection against bacterial challenge between technical replicates within experiments and between experiments, as reflected in the large standard deviations in the relative percent survival (RPS; Table 1). One out of three experiments showed no protection from probiotic treatment. The 24h probiotic pretreatment showed a more consistent level of protection against RE22 challenge (Table 1). In the hatchery trial, larvae treated daily with probiotics for 8 days in the hatchery showed an increase of 28 ± 6 % in relative percent survival as compared to untreated larvae after a laboratory challenge with V. coralliilyticus RE22.

Transcriptome completeness
Depth of sequencing for all the lab transcriptomes was comparable between samples ranging from 16-25M paired end reads whereas HT_RI transcriptomes ranged from 50 -70M reads (Table 2). Sequencing saturation curves for all transcriptomes were close to full saturation, indicating that all but the rarest (least abundant) transcripts would be represented (Figures 1a and b). The alignment rate to the Crassostrea virginica reference genome using HISAT2 ranged from 86 -89% (Table 2).

Differential Expression Analysis
Probiotic treated larval transcriptomes (RI or S4) at each time point (6 or 24 h) were compared to control (0 h) transcriptome for normalization. S4 treated transcriptomes (both 6 h and 24 h) yielded more differentially expressed transcripts when compared to control (0 h) larvae than RI treated larval transcriptomes (p ≤ 0.05) ( Table 3). Larvae treated with probiotics for 24 h yielded more differentially expressed transcripts than larvae treated with probiotics for 6h (Table 3).
Comparison of the number of shared and unique differentially expressed genes across all treatments (Figure 2a) showed a dynamic response to each of the two probiotics.
Overall, larvae treated with S4 for 6 or 24 h have a higher number of differentially expressed transcripts than larvae treated with RI at 6h or 24h. The percentage of DEGs shared between S4 and RI is the same (26%) at 6h or 24h suggesting pronounced effect of treatment as compared to time. Out of the total number of differentially expressed transcripts in response to S4 and RI at 6 and 24h, 50% transcripts were unique to S4 treatment and 21% were unique to RI treatment. Comparison of differentially expressed transcripts in hatchery transcriptomes (HT_RI) (Figure 2b) showed 43% transcripts shared between RI treatments with only 8%, 3% and 8% unique transcripts in RI_5d, RI_12d and RI_16d respectively suggesting more of a treatment effect than time. Refer to supplementary data tables in appendix for descriptions and log fold change values for differentially expressed genes for all comparisons.

GO annotation
A Gene Ontology (GO) term enrichment analysis was performed on all the differentially expressed transcripts in response to probiotic treatment. S4 treatments at both time points shared terms related to recognition and signaling (Figure 3a, 3b). S4 treatment at 6h showed enrichment in "cellular response to stimulus" whereas at 24h it showed enrichment in processes related to activation of receptors and signaling pathways suggesting a progression of immune response to S4. Very few GO terms were significantly enriched among DEGs detected from comparison between the control and larvae exposed to RI in the laboratory and they were mostly related to larval development (not shown). The HT_RI transcriptomes shared enrichment of the term "cytoskeletal organization" (Figures 3c) with the S4 (24h) transcriptomes, but none with the RI laboratory transcriptomes.

KEGG annotation
Consistent with the results of the enrichment analysis, most of the KEGG pathways that were represented by differentially expressed C. virginica larval genes related to signal transduction, immune systems, and endocrine system (Table 4).

Differentially expressed immune genes shared between probiotics
An overview of the immune genes differentially expressed upon exposure to the probiotics is depicted in Figure 4. Transcripts corresponding to the genes for several types of PRRs were modulated by probiotic treatment, out of which Toll-like receptors (TLRs), lectins, recognition protein, peptidoglycan receptor protein (PGRP) and leucine-rich repeat receptors (LRRs) were upregulated, with TLRs and lectins being most upregulated, while scavenger receptors, leucine rich repeat and fibronectin type III domain-containing proteins (LRFN), fibronectin domain containing proteins and C1-q proteins were downregulated. TLR 4, 6 and 13 were consistently upregulated in response to both probiotics with the exception of HT_RI transcriptome where TLR 13 is downregulated (Table 5).
Consistent with the observation that probiotic treatment led to differential expression of several TLR receptors, several transcripts involved in the TLR signaling pathway, including TNF receptor-associated factor 3-like (TRAF3) and mitogen-activated protein kinase kinase kinase 7-like (TAK1), were differentially expressed upon probiotic treatment (Table 6).
Moreover, DEG patterns suggested activation of the NF-kB and MAPK pathways by probiotic exposure. Activation of the NF-kB pathway was indicated by upregulation of activator B-cell lymphoma/leukemia 10-like (BCL10) and downregulation of inhibitor NF-kappa-B inhibitor alpha-like isoform X1 (IkB). Some of the key players of the MAPK pathway including dual specificity mitogen-activated protein kinase kinase 7like (MAP2K7), TAK1, extracellular signal-regulated kinase 2-like (ERK2) were also upregulated in probiotic-treated larvae. Transcripts corresponding to a key molecule of the MAPK pathway, MAP2K7, were uniformly upregulated in almost all probiotic treatments (Table 6).
Probiotic treatment unanimously leads to modulation of three types of major effectors: serine protease inhibitor (SPI), mucin and macrophage-expressed gene 1 protein-like (Mpeg1/Perforin-2) ( Table 7). Serine protease inhibitor Cvspi2 was highly upregulated in all probiotic treatments including HT_RI samples. Digestive cysteine proteinase 2 was highly upregulated in all treatments except HT_RI. Several different types of mucin genes were modulated in larvae due to probiotic treatment. Both secreted gel-forming mucins (MUC2, MUC5A, MUC5B and MUC19) and cell surface mucins (MUC3B, MUC4 and MUC12) were differentially expressed. MUC12 was highly upregulated in almost all probiotic treatments. MUC5AC was highly upregulated in probiotic treatments of 24h and MUC2 was upregulated at 6h. Perforin2 was highly upregulated in all probiotic treated larvae except in HT_RI samples.
Various molecules associated with cytoskeleton reorganization including actin, tubulin, integrin, myosin and septins (Table 8) as well as those related to phagosome, endocytosis, peroxisome and lysosome (Table 10) were differentially expressed in response to probiotics. Prostaglandin G/H synthase 2-like (PTGS2), important in inflammation reaction, was highly upregulated in all but S4(24h) treatment.

Differentially expressed immune genes unique to each probiotic
Transcripts of alpha-1-macroglobulin-like, integrins and antioxidant enzymes were downregulated in larvae exposed to S4 (Table 9). Transcripts corresponding to Tollo (TLR8) and E3 ubiquitin-protein ligase LRSAM1 were highly upregulated in RI(6h) alone. HT_RI transcriptomes showed upregulation of histone H2B-like and GTPase IMAP family member 7-like (GIMAP7) transcripts that were not seen in any other probiotic treatments.

Transcripts involved in antiviral responses
Surprisingly, several genes that are involved in antiviral pathways were also differentially expressed due to probiotic treatment. These included upregulation of recognition receptors (TLR3) for detecting intracellular nucleic acids and transcripts involved in the JAK-STAT and cGAS-STING pathways ( Table 6). Stimulator of interferon genes protein-like (STING), an important part of the cGAS pathway, was upregulated in all probiotic treatments except HT_RI. Interferon induced protein 44 gene was upregulated in the HT_RI sample. E3 ubiquitin-protein ligase TRIM56 was heavily modulated in larvae from both probiotic treatments after a 24h exposure.

Cell death:
Autophagy related ATG9a was highly upregulated in all probiotic treatments except HT_RI (Table 9). Both initiator and executioner caspases in the apoptosis pathway were differentially expressed in probiotic treatments (Table 9). Transcripts for the initiator caspase 2 were upregulated in 6 h treatments while at least one of the executioner caspases 1,3,6 were upregulated in all treatments. Interestingly, caspases 1, 2, 7 and 8 were downregulated and only caspase-14 was upregulated in HT_RI. Several types of baculoviral IAP repeat-containing proteins were differentially expressed in response to probiotic treatments but the type of modulation and type of IAP differed between treatments. Inhibitor of apoptosis was highly up in all probiotic treatments except HT_RI, where GIMAP7 was highly upregulated.

Discussion
Exposure of larvae to probiotics S4 and RI induced the expression of a large variety of immune genes, suggesting a strong immune response comprising of heightened pathogen recognition, activation of immune signaling pathways and production of an arsenal of effectors. This probiotic mechanism of larval immunostimulation is consistent with previous observations that probiotics are cleared from the larvae within 12 -24 h after treatment . These immune effectors activated in larvae upon probiotic exposure may also serve to provide protection against RE22 infection especially in light of the opposite effect of suppression of signaling pathways and lack of crucial effectors seen in response to RE22 challenge (Modak et al., in prep; Ch3 of this dissertation).

Mechanisms shared between probiotics
Overall, the immune response of larvae to each of the probiotics shared many features, including: (a) upregulation of a large variety of pathogen recognition receptors involved in environmental sensing and pathogen detection, followed by (b) activation of multiple signaling pathways; which ultimately led to the production of (c) an arsenal of effectors known to have a role in immune defenses against bacterial pathogens (Figures 4 & 5).
Several probiotics are known to modulate (either activate or suppress) signaling pathways that benefit the host and protect them from pathogens (reviewed in Llewellyn et al., 2017). Usually, probiotics show a very strain specific response (Baarlen et al., 2011, Llewellyn et al., 2017. In this case however, despite the difference in Gram character between S4 and RI, many immune transcripts, especially effectors, were expressed in response to both probiotics.
Overall, differential expression analysis suggests activation of various immune signaling pathways like TLR, NF-kB and MAPK by both probiotics. The TLR pathway is crucial for bivalve innate immune systems. It recognizes a variety of damageassociated molecular patterns (DAMPs) and pathogen-associated molecular patterns (PAMPs) to activate NF-kB and MAPK pathway that protect the host from infection by producing cytokines, chemokines and other effectors (Gerdol et al., 2018). Our findings showing that TLR3, 4, 6, 8 and 13 were upregulated in response to probiotics are consistent with the important role of this pathway in bivalve immune responses, and indicate the potential of probiotics to provide protection against a broad spectrum of pathogens. Such PRR activation by probiotics due to shared cell envelope components like lipopolysaccharides, peptidoglycan, and ß-glucans with pathogens is well known (Pérez-Sánchez et al., 2014). Activation of TLR6 broadens the recognition spectrum to bacteria, fungi, LPS and peptidoglycan (PGN) (Wang et al., 2018). Subsequent activation of the MAPK pathway regulates several important cellular processes like cell proliferation, apoptosis, inflammatory response to pathogens and involved in the innate immunity of oysters (Wang et al., 2018). Activation of host MAPK and NF-kB and other signaling pathways by probiotics is seen in human gut associated probiotics (Thomas & Versalovic et al., 2010, Bermudez-Brito et al., 2012.
This transcriptome analysis also suggests that activation of these pathways leads to increased transcription of a variety of immune effectors. Larvae already equipped with effectors as a result of probiotic treatment can carry out expedited clearing of pathogen upon challenge. Some of these effectors have been shown in previous research to have the potential to be involved in protection against RE22.
Protease inhibitors: All probiotic treatments showed highly upregulated serine protease inhibitor Cvspi2. One of the important virulence factors of RE22 is production of proteases, most notably metalloproteases (Hasegawa et al., 2008), but also potentially serine proteases, which are encoded in the genome , Richards et al., 2018. Presence of serine protease inhibitors might neutralize serine protease attack by RE22 in probiotic pretreated larvae thus playing a significant role in their survival from RE22 infection. cvSI-1 has been shown to play an important role in host defense against Perkinsus marinus by inhibiting proliferation of the parasite (LaPeyre et al., 2009 and is also upregulated in resistant oysters in response to challenge with the pathogen Aliiroseovarious crassostreae  in C. virginica.
Mucins: Mucus is an important line of defense and plays multiple roles in the hostmicrobe interaction . Both secreted gel forming mucins and cell surface mucins modulated by both probiotics work in concert to clear infection (Linden et al., 2008). Both Gram negative and Gram-positive bacteria have been shown to upregulate mucins in humans (Dohrman et al., 1998) which explains how both probiotics could influence their production. Increased production of mucus could buffer action of proteases (Yan et al., 2017) used by pathogenic Vibrio spp. to penetrate mucus and spread infection (Silva et al., 2003). Probiotics modulate the mucus barrier to aid their adhesion thereby preventing invasion of pathogens (Tuomola et al., 1999, Allam and. In addition, oysters can also benefit from presence of vast array of immune recognition and effector proteins in the mucus . Hence, modulation of mucins can have multiple advantages for probiotic pretreated larvae.
Perforin-2 is an important ancient innate immune system effector present in vertebrates as well as invertebrates that functions by forming pores in intracellular and extracellular pathogenic bacteria (McCormack and Podack, 2015). In invertebrates, LPS exposure significantly upregulated a homologue of perforin-2 in a sponge Suberites domuncula (Wiens et al., 2005) and in disk abalone Haliotis discus discus post V. parahemolyticus challenge (Bathige et al., 2014). In C. gigas, Cg-Mpeg1 showed significant antibacterial activity to both Gram-negative and positive bacteria and its transcription level was significantly up-regulated following infection with V. alginolyticus .
Thus, elevated activation of perforin-2 in probiotic pretreated C. virginica larvae might act as an efficient effector against RE22 upon challenge.
Cytoskeletal rearrangements can help in bacterial sensing, compartmentalization of pathogens (Mostowy & Cossart, 2011), autophagy and apoptosis for host protection (Mostowy and Shenoy, 2015) as well as phagocytosis (Vicente-Manzanares and Sánchez-Madrid., 2004). PTGS2, which was upregulated in almost all probiotic treatments, is a key enzyme producing inflammatory prostaglandins and generation of inflammatory response activating the immune system in advance.

Mechanisms unique for each probiotic
Some unique aspects of the probiotic specific response are discussed below:

Specific response to S4:
In addition to protease inhibition, alpha-1-macroglobulin (which was downregulated only in S4) is also involved in complement and coagulation cascades (Xiao et al., 2000) suggesting possible modulation of complement cascades by S4. Integrins (also downregulated in S4(24h)) have been shown to be used by V. splendidus to enter hemocytes and evade immunity (Duperthuy, M, et al., 2011). Antioxidant enzymes were mostly downregulated with S4 treatment, suggesting that S4 treatment does not lead to oxidative damage, unlike pathogenic exposure (Lorgeril et al., 2008.

Specific response to RI:
Tollo (TLR8, upregulated in response to RI) is related to larval innate immune response to Gram negative and positive bacteria and shown to regulate antimicrobial production in Drosophila melanogaster (Akhouayri et al., 2011). E3 ubiquitin-protein ligase LRSAM1 (highly upregulated in RI (6 h)) is a bacterial recognition protein and ubiquitin ligase that defends the cytoplasm from invasive pathogens. It is important for ubiquitindependent autophagy against invading intracellular bacterial pathogens (Huett et al., 2012).
Two unique aspects about HT_RI transcriptome were upregulated transcripts identified as histone H2B-like and GIMAP7. Histones show antimicrobial action against Gram negative bacteria like Escherichia coli (Kawasaki et al., 2008) and in C. gigas has been demonstrated to surround and engulf vibrios (Nikapitiya et al., 2013, Poirier et al., 2014. GIMAP7 is member of GTPase of the immune-associated proteins family that acts an apoptosis regulator (Nitta and Takahama, 2007) and its upregulation suggests inhibition of apoptosis.

Unexpected responses to probiotics
Interestingly, multiple members of antiviral pathways were also modulated in response to probiotics. STING, an important part of the cGAS pathway, was highly upregulated in all lab probiotic treatments. A special STING homolog LvSTING was activated in shrimp in response to V. parahaemolyticus infections that participates in antimicrobial peptide production . Thus, this pathway plays an essential role in host response to pathogen invasion including bacteria, owing to detection of cytosolic DNA recognition and type I IFN production (Tao et al., 2016). Activation of these pathway suggests that probiotic exposure may provide protection against viruses (Thomas et al. 2010, Bermudez-Brito et al., 2012.

Cell death
ATG9a was highly upregulated by all probiotic treatments suggesting activation of autophagy (He and Klionsky, 2009) consistently by both probiotics. Autophagy and septins together restrict cytosolic bacterial replication (Torraca and Mostowy, 2016) and maybe an additional mechanism of action against RE22 invasion.
Various apoptosis inhibitors were highly upregulated in response to both probiotic treatments suggesting overall inhibition of apoptosis in response to probiotics.
However, patterns of expression of apoptotic genes vary across different environmental stressors in bivalves suggesting it is a very complex pathway that is still not completely understood (Gerdol et al., 2018). Surviving C. gigas also showed apoptosis inhibition in response to virulent Vibrio sp. . One of the virulence factors of RE22 is production of hemolysins  showing toxic effects on hemocytes . Inhibition of apoptosis by probiotic pretreatment might result in a higher number of hemocytes (Lee et al., 1993) that can potentially counter the effect of hemolysins secreted by RE22 upon challenge.

Length of probiotic pretreatment for effective protection from challenge
As seen in the results (Table 1), shorter probiotic pretreatment provides variable protection whereas longer pretreatment provides consistent protection from challenge.
Comparison of 6h and 24h transcriptomes showed same key effector mechanisms activated at both time points viz upregulation of serine protease inhibitors, mucins and perforin-2. There are however subtle differences for example in types of PRRs, mucins and septins that are upregulated at 6 h compared to those at 24 h. Certain genes involved in biomineralization and larval development and growth were also upregulated at 24 h.
This supports the observation that longer exposure provides better protection perhaps due to increased pathogen sensing, additional growth effects and longer time for all larvae to respond to probiotic pretreatment. Previous studies have also shown chronic exposure of probiotics work better (Llewellyn et al., 2017).

Conclusion
This study indicates that probiotics use immunomodulation as a mechanism of action that may play a role in the protection conferred against RE22 infection. Although 6 h of pretreatment with probiotics might suffice for some larvae to protect themselves from RE22 challenge, a 24 h pretreatment consistently allows majority of them to elicit the immune responses effective in providing protection. This knowledge might help in designing better management strategies to control larval mortality in hatcheries by use of probiotics as a natural and environmental friendly solution. In the future, it would be beneficial to use this information to target the functional identification of effectors that serve in protecting larvae against RE22 infection.            Figure 4: Overview of the immune responses induced in oyster larvae in response to treatment with probiotics S4 and RI, as measured through high-throughput analysis of differential gene expression. Overall, PRRs including TLRs, lectins, PGRPs and LRRs were upregulated while others were downregulated. Signaling pathways including TLR, NF-kB, MAPK and antiviral pathways including JAK-STAT, cGAS-STING were activated. Immune effectors were activated including mucins, protease inhibitor and perforin-2. Autophagy was activated and apoptosis was inhibited. Antioxidant enzymes were downregulated. Cytoskeleton related molecules including septins were modulated by both probiotics.

Figure 5:
Hypothesized role of selected effectors of immunity whose expression was found to be upregulated in larval oysters in response to probiotic treatment on providing protection against challenge to V. coralliilyticus RE22. Mucin and protease inhibitors provide protection outside the oyster body and perforin-2 providing protection once the pathogen is within oyster tissues. Table 5: Patterns of differential gene expression of immune receptors in oyster larvae in response to probiotic treatment (p ≤ 0.05, upregulation: log fold change ≥ 2, downregulation: log fold change ≤ -2). Yellow denotes downregulation, orange denotes up and downregulation of transcripts mapped to the same gene, red denotes upregulation. HT-RI: larvae treated daily with probiotic Bacillus pumilus RI0695 (RI) for 5, 12 or 16 days; RI_6h: Larvae exposed to RI for 6h; RI_24h: Larvae exposed to RI for 24h; S4_6h: Larvae exposed to S4 for 6h; S4_24h: Larvae exposed to S4 for 24h.  Table 6: Patterns of differential gene expression of immune signaling pathways in oyster larvae in response to probiotic treatment (p ≤ 0.05, upregulation: log fold change ≥ 2, downregulation: log fold change ≤ -2). Yellow denotes downregulation, orange denotes up and downregulation of transcripts mapped to the same gene, red denotes upregulation. HT-RI: larvae treated daily with probiotic Bacillus pumilus RI0695 (RI) for 5, 12 or 16 days; RI_6h: Larvae exposed to RI for 6h; RI_24h: Larvae exposed to RI for 24h; S4_6h: Larvae exposed to S4 for 6h; S4_24h: Larvae exposed to S4 for 24h.

TLR pathway
TNF receptor-associated factor 3-like isoform X3 (TRAF3) tumor necrosis factor receptor superfamily member 1B-like isoform X1 tumor necrosis factor receptor superfamily member mitogen-activated protein kinase kinase kinase 7-like isoform X3 (TAK1)  Table 7: Patterns of differential gene expression of immune effectors in response to probiotic treatment (p ≤ 0.05, upregulation: log fold change ≥ 2, downregulation: log fold change ≤ -2). Yellow denotes downregulation, orange denotes up and downregulation of transcripts mapped to the same gene, red denotes upregulation. HT-RI: larvae treated daily with probiotic Bacillus pumilus RI0695 (RI) for 5, 12 or 16 days; RI_6h: Larvae exposed to RI for 6h; RI_24h: Larvae exposed to RI for 24h; S4_6h: Larvae exposed to S4 for 6h; S4_24h: Larvae exposed to S4 for 24h.  Table 8: Patterns of differential gene expression that are part of cytoskeletal reorganization in oyster larvae in response to probiotic treatment (p ≤ 0.05, upregulation: log fold change ≥ 2, downregulation: log fold change ≤ -2). Yellow denotes downregulation, orange denotes up and downregulation of transcripts mapped to the same gene, red denotes upregulation. HT-RI: larvae treated daily with probiotic Bacillus pumilus RI0695 (RI) for 5, 12 or 16 days; RI_6h: Larvae exposed to RI for 6h; RI_24h: Larvae exposed to RI for 24h; S4_6h: Larvae exposed to S4 for 6h; S4_24h: Larvae exposed to S4 for 24h.  Table 9: Patterns of differential gene expression that are part of apoptosis and autophagy in oyster larvae in response to probiotic treatment (p ≤ 0.05, upregulation: log fold change ≥ 2, downregulation: log fold change ≤ -2). Yellow denotes downregulation, orange denotes up and downregulation of transcripts mapped to the same gene, red denotes upregulation. HT-RI: larvae treated daily with probiotic Bacillus pumilus RI0695 (RI) for 5, 12 or 16 days; RI_6h: Larvae exposed to RI for 6h; RI_24h: Larvae exposed to RI for 24h; S4_6h: Larvae exposed to S4 for 6h; S4_24h: Larvae exposed to S4 for 24h.

Apoptosis
Caspase 1  days; RI_6h: Larvae exposed to RI for 6h; RI_24h: Larvae exposed to RI for 24h; S4_6h: Larvae exposed to S4 for 6h; S4_24h: Larvae exposed to S4 for 24h.  Table 11: Patterns of differential gene expression that are part of metabolism, biomineralization and other processes in oyster larvae in response to probiotic treatment (p ≤ 0.05, upregulation: log fold change ≥ 2, downregulation: log fold change ≤ -2). Yellow denotes downregulation, orange denotes up and downregulation of transcripts mapped to the same gene, red denotes upregulation. HT-RI: larvae treated daily with probiotic Bacillus pumilus RI0695 (RI) for 5, 12 or 16 days; RI_6h: Larvae exposed to RI for 6h; RI_24h: Larvae exposed to RI for 24h; S4_6h: Larvae exposed to S4 for 6h; S4_24h: Larvae exposed to S4 for 24h.

HT_RI RI_6h RI_24h S4_6h S4_24h
Others furin-like protease kpc-1 isoform X1 can be safely and effectively used to limit negative impacts of vibriosis in shellfish hatcheries. Understanding host-microbe interactions between C. virginica larvae and pathogen or between larvae and probiotics would immensely help in designing protocols of probiotic use commercially.
This research showed the swift progression of disease both in terms of rapidly increasing mortality post 14h of exposure as well as impact on host immune system.
Immunological responses of C. virginica larvae to pathogen V. coralliilyticus RE22, as measured through transcriptome analysis, suggest the ability of vibrio exposure to suppress immune-related pathway activation and immune effector production. The research also highlights the need and suitability of preventative measures like probiotics rather than treatment options to protect larvae from effects of V. coralliilyticus RE22.
This dissertation research on the immunological responses of C. virginica larvae to both probiotics B. pumilus RI06-95 and P. inhibens S4 shows that the immunosuppression by RE22 may be counteracted by probiotics 'priming' of the larval immune response. This research demonstrates the ability of both probiotics to activate pathogen recognition receptors (PRRs) that could aid in pathogen detection, activation of immune signaling pathways and production of immune effectors that could potentially aid in inactivation of RE22 and its virulence factors.
A hypothesized model based on the findings of this dissertation research and previously published work is proposed here (Fig 1). When C. virginica larvae are pretreated with probiotics, RI and S4 for 6 to 24 h, most larvae are protected from RE22 challenge (Fig1-1). A more prolonged 24 h pretreatment (versus 6 h) allows for more consistent elicitation of immune responses, and therefore more consistent levels of protection against RE22. Immune responses include activation of PRRs, immune signaling pathways and production of immune effectors like mucins, serine protease inhibitors and perforn-2 (Fig1-2). Oysters have a high basal rate of apoptosis that regulate hemocyte number (Sokolova 2009). Transcriptomic data suggests treatment with probiotics may inhibit hemocyte apoptosis, leading to increase in the number of hemocytes (Fig1-3). This immunostimulation likely contributes to clearing probiotics from the system , but also may contribute to counteracting RE22 virulence. When probiotic pretreated (and hence immunostimulated larvae) are challenged with RE22, a series of changes brought about by the probiotics in the host may assist the larvae in blocking RE22 (Fig1-4). Increased mucin production may enhance the epithelial barrier blocking penetration and prevent adhesion of pathogen.
Increased production of serine protease inhibitors may help to counter the effect of serine proteases potentially produced by RE22. This immunomodulation would complement other mechanisms of action of probiotics. Probiotic biofilm established during the pretreatment period may reduce colonization sites for RE22 competitively excluding them from colonizing the gut. Biofilm formation and competition assays between S4 and RE22 showed pretreatment with S4 excludes RE22 .
The draft genome of RI suggested its ability to form biofilms  but there is no experimental data to support it yet. Antibiotic tropodithietic acid (TDA) produced by S4 also aids in eliminating RE22 . S4 also secretes Nacyl homoserine lactones (AHLs) that quorum quench RE22 metalloprotease gene expression that are a crucial part of its virulence (Zhao et al., 2018). Increased production of perforin-2 due to probiotic pretreatment may also aid in neutralizing pathogens both intracellularly and extracellularly within oyster tissues. Increased number of hemocytes owing to apoptosis inhibition post probiotic treatment may increase phagocytic pressure on RE22 as well as buffer cytotoxic effects of hemolysins secreted by RE22 (Fig1-5) that diminish hemocyte survival (Gomez-Leon et al., 2008).
All these effects probably work in concert to allow more probiotic pretreated C.
virginica larvae to survive post RE22 challenge than those without probiotic pretreatment, by effectively reducing the infective dose of RE22 (Fig1-4) and providing larvae with mechanisms to further neutralize and kill RE22 within the oyster tissues (Fig1-5), leading to increased survival (Fig1-6). Due to effective clearing of probiotics within oysters due to the larval immune response, however, their protective effect diminishes over time as also seen in experimental evidence (Karim et al., 2013) unless probiotics are applied repeatedly.
Immune effectors produced in response to probiotics, specifically highlighted in this study are highly suitable in blocking virulence factors and pathogenesis of RE22.
However, application of probiotics and their overall immunostimulatory effect may likely help in protecting larvae from other bacterial and viral infections. Thus, this research advocates use of probiotic formulations in commercial shellfish aquaculture for their beneficial effects. In addition, it provides new insights in oyster immunity in response to non-pathogenic bacteria and the crosstalk between host and probiotics.   Table 2: Differentially expressed genes with log fold change for probiotic or pathogen treatments when compared to control (Con 0 h) in laboratory transcriptomes (p ≤ 0.05, upregulation: log fold change ≥ 2, downregulation: log fold change ≤ -2). RI_6h: Larvae exposed to RI for 6h; RI_24h: Larvae exposed to RI for 24h; S4_6h: Larvae exposed to S4 for 6h; S4_24h: Larvae exposed to S4 for 24h; RE22_6h: Larvae exposed to RE22 for 6h.