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MICROBIAL ECOLOGY OF THE BIVALVIA, WITH AN EMPHASIS ON THE FAMILY OSTREIDAE.

INTRODUCTION

A burgeoning interest in microbial ecology research over the past 20 years is attributed to both rapid improvements in the technology used to assess microbial communities and a decline in associated costs. Next-generation sequencing (NGS) of the 16S rRNA gene has spurred a rapid increase in microbiome research aimed at a broad range of organisms and environments. Whereas major research efforts have focused on humans (e.g., NIH Human Microbiome Project) and direct applications (i.e., health and disease), general ecological studies have also been carried out on invertebrates. Invertebrates are useful both as model organisms and as indicators of ecosystem health; research aimed at understanding the natural composition and variation of invertebrate microbiomes can provide valuable information for downstream questions.

The colonization and maintenance of microbial communities within organisms are not the result of chance interaction. Bacteria, in particular, have formed symbiotic (commensal, mutualistic, and pathogenic) associations on and in the surfaces, tissues, and organs of all eukaryotic organisms. These interactions have influenced the evolution of prokaryotes and eukaryotes and, in many cases, have led to mutually beneficial and even obligatory relationships (Round & Mazmanian 2009, Fraune et al. 2010, Lee & Mazmanian 2010, Nyholm & Graf 2012, Buffie & Pamer 2013, Mendes et al. 2013, Huang et al. 2014). Host organisms reap the reward of improved digestion, nutrient absorption, immune system development, and protection from invading pathogens (Crosby et al. 1990, Harris 1993, Wold & Alderberth 2000, Guarner & Malagelada 2003, Ley et al. 2005, Turnbaugh et al. 2007, Mazmanian et al. 2008, Kau et al. 2011, Forberg et al. 2012, Dishaw et al. 2014a). In turn, microbes receive a stable environment and a constant nutrient supply (Karasov & Martinez del Rio 2007, Ley et al. 2008a, Neish 2009). It should be noted, however, that many uncertainties still remain as to whether hosts and their microbes, or "hologenomes," evolve as a unit, or separately (Moran & Sloan 2015). Regardless, host-associated microbiota are an integral component in the development and functioning of their hosts.

Some of these host-prokaryote relationships are very specific. Shipworms, a species of marine bivalve molluscs, harbor in their gills a community of Gammaproteobacteria which aid in the digestion of cellulose, as wood comprises the entirety of their diet. These bacteria are found across many shipworm species distributed worldwide (Distel et al. 1991, Distel et al. 2002). The Hawaiian Bobtail squid Euprymna scolopes Berry (1913) harbors the bacterium Vibrio fischeri (Beijerinck, 1889) in a light organ to avoid predators by counter-illumination when hunting at night (Jones & Nishiguchi 2004, Nyholm & McFall-Ngai 2004). Newly hatched squid acquire the bacterium from the environment, selecting V. fischeri over all other bacterial species (reviewed in Nyholm & McFall-Ngai 2004). The specificity is maintained by a number of host and symbiont factors.

Less specific interactions have also been reported, and interest in the ecology of host-associated microbiota has increased in recent years (Evans et al. 2013, Mavrommatis et al. 2013, Mendes et al. 2013). Research on this topic has included a number of species, terrestrial and aquatic, ranging from sponges to humans. For example, studies on sponges, coral, and shellfish have shown that microbial communities found in the host can be distinct from habitat-specific bacteria found in the environment (Harris 1993, Rosenberg et al. 2007, Fan et al. 2012, Schmitt et al. 2012, Kimes et al. 2013). In humans, tissue-specific communities have been identified within individuals (Turnbaugh et al. 2007, Human Microbiome Project Consortium 2012). In addition, core microbiomes, in which the microbiota are conserved across individuals of the same species, have been discovered for a number of hosts (zebrafish--Roeselers et al. 2011; tunicate--Dishaw et al. 2014b; shrimp--Rungrassamee et al. 2014). This phenomenon may indicate a specific functional role of the conserved bacteria.

Interest in the microbial communities of bivalves has a long history in biology. Scientists have investigated the prokaryotes associated with oysters for over 100 years (Round 1914, Colwell& Sparks 1967, Kueh & Chan 1985, DePaola et al. 1990, Green & Barnes 2010, Lokmer & Wegner 2015). Bivalves, especially oysters, are of particular interest as a result of their importance in ecosystem functioning and as an economic commodity for humans. For example, the eastern oyster [Crassostrea virginica (Gmelin, 1791)] fishery is valued at >$196 million annually in the United States alone (NMFS 2015). Consequently, understanding the factors that mitigate the health of bivalves has implications for the environment and the global economy. Because of their economic value and consumption by humans, the vast majority of the literature focuses on disease and zoonotic pathogens. The relationship between bivalves, their commensal microbiota, and the environment has traditionally been less well studied and understood. Over three decades ago, Harris (1993) summarized the literature pertaining to the gut microbial communities of aquatic invertebrates. Although important in its time, an updated review that focuses solely on the Bivalvia and includes more relevant culture-independent studies and data on the microbial communities of tissues other than the gut is needed.

This review focuses on research related to the ecology of bivalve-associated microbial communities and less on indicator organisms and pathogens. Specifically, their composition and structure, spatial and temporal distribution, host specificity, and relationship to bivalve physiological functioning are summarized. Most of the recently published research has focused on the feeding and digestive organs (i.e., gill and gut) and the hemolymph. Therefore, the digestive and circulatory systems are highlighted in this review. Research on other aquatic invertebrates is used as a context for the work that has been carried out on bivalves. Finally, the importance of these studies in the larger field of microbial ecology and future research directions are discussed.

MICROBIAL COMMUNITIES OF AQUATIC INVERTEBRATES

In the aquatic environment, differences in bacterial composition between seawater and hosts indicate a niche selection by bacteria in which the host represents a favorable environment (La Valley et al. 2009, Lokmer & Wegner 2015). Interspecific differences in the microbiota at one location, and intraspecific similarities in the microbiota of geographically separate individuals, indicate potential species-specific relationships (Fraune & Bosch 2007, Zurel et al. 2011, Dishaw et al. 2012, Trabal et al. 2012, Dishaw et al. 2014b, Pierce et al. 2016). Although some studies have demonstrated such specificity, others have found diverse and variable microbiota within a species and site (e.g., amphipods--Mengoni et al. 2013) and differences within a species between sites (e.g., Crassostrea spp. oysters--King et al. 2012, Trabal et al. 2014; queen conch--Carrascal et al. 2014; Brachidontes mussel--Cleary et al. 2015). The influence of stochastic processes may also be a factor for diversity, which would help explain variation in the microbiome among organisms separated geographically. These observed differences may be attributed to chance host-bacterial interactions (Lankau et al. 2012).

A few studies have postulated that the host itself imparts control on the bacterial communities it maintains, which may help to explain the widespread trend of core microbiomes across diverse phylogenies. One striking example of this concept was seen in research with the hydrozoan Hydra, where environmental and laboratory-reared organisms harbored the same microbiota (Fraune & Bosch 2007). In addition. Hydra oligactis Pallas, 1766 and Hydra vulgaris Pallas, 1766 reared under identical conditions maintained distinct communities (Fraune & Bosch 2010). Host genetic factors may be a driving force in colonization control (Dishaw et al. 2014a). It is likely, however, that both extrinsic (environmental) and intrinsic (host) factors are at play in shaping and mediating the pro-karyotic communities of aquatic organisms.

To understand the microbiomes of marine bivalve molluscs, research on more well-studied aquatic suspension feeders provides a relevant analog. Marine sponges, although they do not contain a gut, are important marine suspension feeders that harbor bacteria in high densities of up to [10.sup.9] cells/[cm.sup.3] of tissue (Hentschel et al. 2012). They contain microbiota distinct from the seawater, although it is thought that many sponges do recruit symbionts from the surrounding water (Fan et al. 2012). A high diversity of bacteria is found, with nearly as many operational taxonomic units (OTUs) recovered in sponges as in seawater samples, and more than 32 phyla regularly associated with many sponge species (Reveillaud et al. 2014, Thomas et al. 2016). These bacterial assemblages are highly conserved among poriferan species, over both time and space, and conservation is thought to be due in part to the microbes fulfilling specific functional roles for the sponges (Fan et al. 2012, Trindade-Silva et al. 2012, Blanquer et al. 2013, Reveillaud et al. 2014). Part of the conservation among species can be linked to the vertical transmission of bacteria from the parental tissue, a result of reproduction by fragmentation (Webster et al. 2009, Hentschel et al. 2012). Despite the differences in life history, structure, and reproduction, sponges provide a simplistic model to relate to other suspension feeders.

Like sponges, corals have been found to harbor bacterial communities that differ from the surrounding environment. Within individual soft and hard corals, bacteria inhabiting the mucus layer covering the colony differ from those within the skeleton and tissue, indicating organ specificity and preference (Rosenberg et al. 2007). The mucus layer serves as a nutrientrich environment that supports a diverse and species-specific bacterial community (Meikle et al. 1998, Rohwer et al. 2001, Rohwer et al. 2002). The microbiota benefit the coral by absorbing UV radiation, supplying nutrients, and protecting it from pathogens (Anthony & Fabricius 2000, Shnit-Orland & Kushmaro 2009, Ravindran et al. 2013). Under stressful conditions, however, these bacteria can become pathogenic (Lesser et al. 2007, Thurber et al. 2009). Research has shown more definitive differences among healthy, diseased, and dead coral microbiomes, which are defined by health status regardless of species or geography (Frias-Lopez et al. 2002, Roder et al. 2014).

As with other suspension feeders, species-specific microbial communities have been observed in tunicates (Dishaw et al. 2014b). This trend has been consistent formultiple species, even when tunicates were separated geographically with little genetic flow (Cahill et al. 2016). Tissue-specific microbial communities are also likely, with the gut and tunic microbiomes distinct from one another and from environmental samples (Blasiak et al. 2014, Cahill et al. 2016). By contrast to sponges, the core microbiome of tunicates comprises a smaller number of phyla and OTUs (Dishaw et al. 2014b, Cahill et al. 2016).

These and other host-microbiome studies have focused largely on the diversity of microbial communities, with an emphasis on their spatial and temporal distributions as well as their relationship to the environment, and thus the surrounding seawater. As sponges, corals, and tunicates are also suspension-feeding invertebrates, the results of these studies provide a useful baseline for comparison with the microbial communities of bivalves. When considering the host microbiome, both functional and genetic diversity of the community should be evaluated. The microbiome can be divided into two components: the core, or resident community, and the transient community. The transient microbial community may be more seasonally affected than the core, attributed to the passing of seawater and food through the host. The functional and genetic diversity of the core and transient microbiomes are influenced by numerous extrinsic and intrinsic factors (Fig. 1).

BACTERIAL ASSOCIATIONS WITH SUSPENSION-FEEDING BIVALVE MOLLUSCS

Suspension feeders, such as oysters and mussels, often dominate the macrobenthos in coastal systems and play a major role in ecosystem processes. Bivalves interact with the water column by cycling dissolved nutrients, removing plankton and aggregated material, depositing feces and pseudofeces, and contributing to the concentration of transparent exopolymer particles (Pile et al. 1997, Prins et al. 1998, Newell 2004, Heinonen et al. 2007, Li et al. 2008, Dame 1993). Because of their dynamic interactions with the water column, encounter rates with microbes are high. Bivalves have the ability to filter large quantities of water (e.g., 3-9 L/h/g dry mass for oysters; Newell et al. 2005, Cranford et al. 2011), and so come into contact with, and pass through their bodies, both free-living and particle-associated microbes. The bacteria with which they interact and harbor have the potential to be particularly important to physiological and biochemical enantiostasis.

Microbes associated with bivalves have been well studied for more than a century (i.e., Round 1914). These studies, however, largely focused on the uptake and depuration of bacteria pathogenic to humans (e.g., Vibrio spp.; DePaola et al. 1990, Froelich & Oliver 2013), or on indicators of such pathogens [e.g., Escherichia coli (Migula, 1895) Colwell & Liston 1960, La Valley et al. 2009]. In addition, in early studies, homogenization of whole animals was routine and evaluation of the microbiota was limited by the sole use of culture-dependent media (Murchelano & Brown 1968, Kueh & Chan 1985). Broad-based ecological studies have only become possible with the recent development of molecular techniques (e.g., amplicon sequencing), including research on tissue-specific communities (Hernandez-Zarate & Olmos-Soto 2006, Green & Barnes 2010, Meisterhans et al. 2016, Pierce et al. 2016).

Research using culture-dependent methods demonstrated that Achromobacter, Aeromonas, Altermonas, Pseudomonas, Flavobacterium/Cytophaga, Micrococcus, and Vibrio were bacterial genera commonly isolated from bivalves (Colwell & Liston 1960, Vasconcelos & Lee 1972, Pillai 1980, Kueh & Chan 1985, Olafsen et al. 1993, Pujalte et al. 1999). Findings also indicated that bacterial concentrations in bivalves are higher by an order of magnitude than those in seawater, and dominance of culturable bacteria is classically linked to seasonality and water temperature (Motes et al. 1998, Cavallo et al. 2009, Zurel et al. 2011). More recent culture-independent work has shown that although these genera are commonly present, they do not necessarily represent most of the community (Romero et al. 2002, Winters et al. 2010, Wegner et al. 2013, Trabal et al. 2014, Li & Wang 2017, Pierce & Ward in review).

As with other organisms (e.g., corals and humans), tissue-and location-specific microbial communities have been identified in bivalves. Parts of the digestive system tend to harbor the highest concentrations of bacteria in the animal, the most notable areas being the stomach, gastric juices, crystalline style, and digestive diverticula (Kueh & Chan 1985). Differences between stomach and hind gut microbiomes have been observed in Crassostrea virginica, with the stomach having a lower diversity of phyla present and a core microbiome with fewer OTUs (King et al. 2012). Using fluorescence in situ hybridization, Hernandez-Zarate and Olmos-Soto (2006) found that the gills of oysters contained a higher diversity of bacteria than either digestive glands or gonads. Total bacterial counts comparable to seawater have also been found in the gills, mantle fluid, and adductor muscles of bivalves (Kueh & Chan 1985), potentially as a result of the mantle cavity and associated structures being regularly bathed in seawater during feeding. Like the gills and pallial fluid, bivalve hemolymph is in contact with the environment, and thus may harbor a more diverse microbial community compared with the organs. Lokmer et al. (2016b) found high diversity in the hemolymph of Crassostrea gigas (Thunberg, 1793) compared with the gut, gill, and mantle tissues. These differences in diversity may be attributable to specialized roles of bacteria associated with specific tissues (Lokmer et al. 2016b, Pierce et al. 2016).

THE BIVALVE MICROBIOME

Community Composition

Results from studies employing molecular methodologies, including NGS, have found that Proteobacteria and Cyanobacteria are abundant members of bivalve gut communities, regardless of host species (King et al. 2012, Trabal et al. 2014, Pierce & Ward in review). Cyanobacteria, however, are likely derived from the particle diet, and not long-term residents of the gut community (Pierce 2016). Trabal et al. (2014) depurated oysters before microbial community characterization and did not observe Cyanobacteria represented in any of the three Crassostrea spp. analyzed. In individual oysters, Proteobacteria may account for >50% of the total abundance, with Alpha (a)-and Gamma (y)-proteobacteria consistently reported as the most abundant (Pujalte et al. 1999, Hernandez-Zarate & Olmos-Soto 2006, Green & Barnes 2010, Fernandez-Piquer et al. 2012, Trabal et al. 2014, Wang et al. 2016, Lokmer et al. 2016b, Rong et al. 2018, Pierce & Ward in review). Within the y-Proteobacteria, Shewanella, Pseudoaltermonas, Oceanospiralles, and Vibrio are major taxa reported. Results from research on mussels mirror the trends found in oysters with regard to the abundance of Proteobacteria and its classes (Pfister et al. 2010, Winters et al. 2011, Cleary et al. 2015, Cleary & Polonia 2018, Vezzulli et al. 2018). Other dominant phyla identified in the gut of oysters, mussels, and clams include Tenericutes, Bacteroidetes, Actino-bacteria, Firmicutes, Chlamydiae, Fusobacteria, Spirochaetes, Chloroflexi, Plantomycetes, and Verrucomicrobia (Fernandez-Piquer et al. 2012, King et al. 2012, Trabal et al. 2012, Trabal et al. 2014, Cleary et al. 2015, Roterman et al. 2015, Arfken et al. 2017, Pierce & Ward in review). Operational taxonomic units from the genus Mycoplasma are also consistently found associated with bivalves, often in high abundances (Winters et al. 2011, King et al. 2012, Cleary et al. 2015, Lokmer et al. 2016b, Arfken et al. 2017, Milan et al. 2018, Pierce & Ward in review). In particular, Crassostrea spp., have been observed to harbor high abundances of Mycoplasma. Another notable group are the sulfate-reducing bacteria, with sulfur metabolizers from the Deltaproteobacteria found in mussels and oysters (Vezzulli et al. 2018, Pierce & Ward in review).

Like the gut, gill communities of Pacific oysters Crassostrea gigas, also are dominated by Proteobacteria, with most OTUs belonging to the [alpha]-proteobacteria class (Wegner et al. 2013). Hernandez-Zarate and Olmos-Soto (2006) observed that C. gigas gills were colonized by Bacteroidetes and [gamma]-Proteobacteria. Bivalves from the genus Chama have been observed to contain mostly the [gamma]-Proteobacteria family Oceanospirillales in the gill tissue (Zurel et al. 2011).

Healthy Crassostrea gigas hemolymph is dominated by Flavobacteria and a-, [gamma]-, and Epsilon (t)-proteobacteria (Lokmer & Wegner 2015, Lokmer et al. 2016b), whereas pathogenic Arcobacter was associated with moribund oyster hemolymph (Lokmer & Wegner 2015). Vezzulli et al. (2018) found the y-proteobacterium Pseudoaltermonas in C. gigas and Mytilus galloprovincialis Lamarck (1819) hemolymph. Similarly, y- and Z-proteobacteria were components in Crassostrea angulata hemolymph, with Pseudoalteromonas and Arcobacter genera representing the two Proteobacteria classes, respectively (Zhang et al. 2018). The ability of opportunistic pathogens to survive the antimicrobial components of the hemolymph could be a result of their own production of antimicrobial compounds (Vezzulli et al. 2018).

Spatial, Temporal, and Tissue-Specific Trends

The seasonal variability of bivalve microbial communities demonstrates a direct relationship between temperature and bacterial abundance, diversity, and structure (Motes et al. 1998, Pujalte et al. 1999, Cavallo et al. 2009, Zurel et al. 2011, Roterman et al. 2015, Pierce et al. 2016, Wang et al. 2016, Lokmer et al. 2016a, Pierce & Ward in review). As many bacteria experience optimal growth at higher water temperature ranges, a decrease in bacterial counts, as well as diversity, in cold, winter months is not unexpected. With some variation, these trends hold true, regardless of bivalve host species or tissue type.

In a seasonal study across three sites in Long Island Sound, changes in the microbial composition and functional diversity of the gut and pallial fluid of Crassostrea virginica were correlated with temperature, regardless of site (Pierce et al. 2016). The microbial communities of oysters from all sites were most similar across months that had similar seawater temperatures, with warm months (>20[degrees]C) distinctly different from cold months (<10[degrees]C) (Pierce et al. 2016). Results of a second seasonal study again showed that gut microbial communities of eastern oysters in Long Island Sound grouped by month (temperature; Pierce & Ward in review). Functional diversity of microbial communities was also directly correlated with temperature, with richness of carbon sources used the highest in summer months, moderate in fall months, and lowest in winter months (Pierce et al. 2016, Pierce & Ward in review). At the phylum level, bacterial community structure remained stable seasonally, with shifts seen in abundance but not diversity. At the OTU level, the same trend was observed, with similar Shannon diversity (H') values throughout the summer and fall. In another study, microbial communities in the gut of Crassostrea hongkongensis Lam & B. Morton (2003) similarly maintained highest H' values from May to November, with a significant decrease in winter months (Wang et al. 2016). Higher microbial species richness in summer (August) versus winter (March) was seen in the gills of Chama pacifica Broderip (1835; eastern Mediterranean Sea; Zurel et al. 2011). In contrast with C. pacifica, however, Chama savignvi Jousseaume in Lamy (1921; Northern Red Sea) maintained more consistent species richness across the year (Zurel et al. 2011). Similar to the finding with Chama spp. from the same locations, gill-associated microbial communities of Spondylus spp. had significantly higher diversity indices in months with warmer water temperatures than gill communities sampled during low water temperatures (Roterman et al. 2015). It is important to note that for these two studies on bivalve gills, water temperatures varied only between 21[degrees]C-28[degrees]C (Mediterranean) or 16[degrees]C-31[degrees]C (Red Sea). Lokmer et al. (2016a) also observed a positive correlation between sea-water temperature and the alpha diversity of the hemolymph microbiome of Crassostrea gigas at a single site in the Wadden Sea.

In a temporal comparison of bivalve species, blue mussels (Mytilus edulis Linnaeus, 1758) harbored gut microbial communities that were more taxonomically similar to one another than they were to eastern oysters. Like oysters, mussels also maintained high genetic diversity over the summer and fall, with the diversity and abundance of the gut microbiota decreasing during the winter (Pierce & Ward in review). Unlike oysters, the microbiota in the gut of blue mussels maintained high functional diversity, richness, and evenness seasonally (Pierce & Ward in review). This functional stability of the microbial communities in the gut of mussels may be attributed to host physiological activity. By contrast to oysters, mussels remain active during winter months with clearance rates (L/g/h) that can be similar across temperature ranges of 0.5[degrees]C-12[degrees]C (Cranford et al. 2011). Vezzulli et al. (2018) also compared the gut and hemolymph of mussels (Mytilus galloprovincialis) and oysters (Crassostrea gigas) at a single site and found that both bivalves maintained distinct gut microbial communities, with some overlap, but hemolymph communities were largely similar. Again, this may be indicative of the pallial cavity's contact with seawater.

Temporally, samples from Pierce et al. (2016) and Pierce and Ward (in review) clustered together giving further indication that oyster gut communities are stable not only spatially within Long Island Sound but also across years during the same season. The overall similarity of microbial communities across multiple years could be attributed to high gene flow between oysters at different sites within Long Island Sound (Hedgecock 1994). Other workers, however, have observed species-specific microbial communities between geographically disparate sites, with no potential gene flow. In a comparison of two species of jewel box clams from the genus Chama, both species harbored similar gill-associated microbial communities during summer months, despite being collected from two geographically separate water bodies (Zurel et al. 2011). Thorny oysters from the eastern Mediterranean and the Northern Red Sea, which are more closely related to scallops than to true oysters, were also evaluated for their gill microbiota and displayed species-specific trends, regardless of geography (Roterman et al. 2015). The Spondylus spp. evaluated by Roterman et al. (2015) harbored gill microbial communities which were more similar to their host species at different locations than to other host species within the genus at the same location, reinforcing the idea of species-specific communities, with the host being a better predictor than environment for bacterial community structure in the gill.

By contrast, some studies on the gut microbiota of oysters show a different trend, demonstrating intraspecific variation between sites (King et al. 2012, Trabal et al. 2012, Trabal et al. 2014). Trabal et al. 2014 found that adult Crassostrea corteziensis (Hertlein, 1951) and Crassostrea sikamea (Amemiya, 1928) from the same grow-out site harbored gut microbial communities that were more similar to each other than to their respective species at another site. Meanwhile, Crassostrea gigas from different grow-out sites maintained high intraspecific variation in their gut microbiomes, even within a site (Trabal et al. 2012, Trabal et al. 2014), making definitive trends difficult to interpret. Similarly, Lokmer et al. (2016a) found small-scale variability in the C. gigas hemolymph microbiome both within individuals and within a site.

Other studies on bivalves also report intraspecific differences in the microbiome between sites. The gill- and gut-associated microbiota of Manila clams [Ruditapes philippinarum (Adams & Reeve, 1850)] were compared between a marine and a brackish habitat within Arcachon Bay on the French Atlantic coast. Significant differences in the microbiota were found between the gill and gut, and between site, although they did maintain some overlap in community structure at the two sites, with some ANOSIM tests being only marginally significant (Meisterhans et al. 2016). In a study comparing mussels from isolated marine lakes and open marine waters, habitat was found to be a significant predictor of bacterial composition of whole mussels (Cleary & Polonia 2018).

On a tissue level, organ-specific microbial communities are a common feature of bivalves. The gill, gut, and hemolymph have been the most well studied sites in oysters, mussels, and clams. Overall, in oysters, a lower microbial diversity in the gut compared with other organs has been observed (Hernandez-Zarate & Olmos-Soto 2006, Pierce et al. 2016, Lokmer et al. 2016b), indicating specialization. This finding is not surprising, given that microbes which thrive within the gut of organisms must tolerate more extreme conditions (i.e., low pH) (Hood et al. 1971). Variation in functional diversity between sites was greater in gut microbial communities than in pallial fluid microbial communities (Pierce et al. 2016). The functional similarity of pallial fluid communities is likely attributable to its constant contact with the environment, as the water-pumping activity of bivalves baths the pallial cavity in seawater. As a result of this contact, the pallial cavity is likely composed of a broader proportion of transient microbes found in seawater. Hemolymph may exhibit a similar phenomenon, with high diversity reported (Lokmer & Wegner 2015, Lokmer et al. 2016b). Similar to the results of Pierce et al. (2016), Vezzulli et al. (2018) reported that hemolymph microbial communities were more similar in composition between different bivalve species from the same site than gut communities were to one another. Tissue-specific communities were also found in the gut and gills of Manila clams (Meisterhans et al. 2016).

Measures of diversity such as richness and evenness of bivalve-associated communities vary from study to study, depending on a myriad of factors such as tissue type, environmental conditions, and host species and host ontogeny. General trends are apparent, and recent studies using NGS methodologies have been summarized in Table 1. Most NGS studies continue to highlight the prevalence of Gammaproteobacteria from the digestive gland of adult Crassostrea spp., followed by Alpha- and Betaproteobaeteria (Hernandez-Zarate & Olmos-Soto 2006, Trabal et al. 2012, Trabal et al. 2014, Wang et al.

2016, Pierce & Ward in review). The number of OTUs observed (species richness) and Shannon diversity (H') values for these bacterial communities were similar across studies, generally ranging from 200 to 400 OTUs per individual and an alpha diversity of 2-4 (H') (Table 1). These trends seem most consistent across genera within the same tissue types. For example, results from Lokmer et al. (2016b) showed species richness and diversity values in the gut of Crassostrea gigas that were similar to those found in Crassostrea virginica (Pierce & Ward in review). A reduced number of phyla in Trabal et al. (2014) and King et al. (2012) compared with other work could be attributed to the starvation of oysters by Trabal et al. (2014) before analysis and a low sample size (n = 3) used in King et al. (2012). Research on mussels highlights that they tend to have a higher number of OTUs and H' index compared with oysters (Table 1).

It has been hypothesized that the host imparts control over the maintenance of its microbiota (Dishaw et al. 2012), and host genotype has been shown to influence microbiota (Wegner et al. 2013, Lokmer et al. 2016a). Differences in the results of the aforementioned studies may be attributable to differences at the population or species level (e.g., Crassostrea gigas versus Crassostrea virginica, versus Crassostrea corteziensis), and differences in environmental conditions (e.g., salinity, bacterioplankton/diet, temperature, and sediment type). In addition, incongruences among studies may be the result of applied methodologies. For example, the use of different primer sets results in the amplification of different bacterial taxa. The studies described in this section generally analyzed the V1-V2, V3-V5, V4, or V6 hypervariable regions of the 16S gene.

Results of studies examining temporal and spatial changes in the microbiome of other marine invertebrates are similar to those for bivalves. A spatially stable microbiome has been observed in sponges, corals, and tunicates (Fan et al. 2012, Dishaw et al. 2014b, Roder et al. 2014). Many corals and sponges maintain microbial community structures that have been described as species specific (Rohwer et al. 2002, Trindade-Silva et al. 2012). Like bivalves, these organisms are suspension-feeding sessile marine invertebrates. Feeding mechanism may be another factor that contributes to the level and patterns of diversity observed in host-associated microbiomes.

Core Microbiota

Among studies, there are organ-specific shared microbial taxa despite bivalve host species and geographic location. Differences in experimental design makes comparison of microbial communities difficult, but some studies included depuration, to remove transient microbes, allowing a clearer picture of resident microbes.

Seasonally, Crassostrea virginica maintained 108 shared OTUs in their gut microbiomes across season and multiple years (Pierce et al. 2016, Pierce & Ward in review). A core microbiome of 161 OTUs was maintained by Mytilus edulis from all seasons within a year (Pierce & Ward in review). Together, both bivalves possessed a shared core of 117 OTUs or 11%-15% of the total OTUs observed (Pierce & Ward in review). In other marine organisms, fewer core OTUs have been observed (e.g., zebrafish--21 OTUs, Roeselers et al. 2011; shrimp--18 OTUs, Rungrassamee et al. 2014). Differences between organisms could be a result of diet, as herbivorous organisms generally have the highest bacterial diversity in their gut communities (Ley et al. 2008a,b). As diet has been shown to influence the structure of microbial communities in mammals (Ley et al. 2008b), the similarities between these other hosts and bivalves may be of significance. Ley et al. (2008a) found that herbivorous mammals had the highest bacterial diversity at the phylum and OTU level compared with carnivores and omnivores. Most humans, for example, have microbial communities represented by 6 phyla (Ley et al. 2006). Individual adult zebrafish are omnivorous and harbor gut microbial communities with 100-200 OTUs from approximately 20 phyla (Roeselers et al. 2011, Stephens et al. 2016). In keeping with this trend, suspension-feeding marine sponges maintain highly diverse microbial communities, with more than 32 phyla commonly observed from any one species (Reveillaud et al. 2014, Thomas et al. 2016). The high number of core OTUs reported in the literature for herbivorous bivalves compared with carnivorous or omnivorous marine organisms is consistent with previously described influences of diet (Ley et al. 2008a,b).

Trabal et al. (2014) noted that by site, all three species of Crassostrea oysters maintained a core gut microbiome. Post-larval oysters had the highest similarity in their microbiomes, regardless of host species, but the similarity in composition and abundance of their microbes decreased with age and transplantation from cultivation sites to grow-out sites. At the same grow-out site, Crassostrea sikemea and Crassostrea corteziensis maintained a core gut microbiome, including the Betaproteo-bacterium Burkholderia (Trabal et al. 2012, Trabal et al. 2014). Mycoplasma OTUs (class Mollicutes) were one of the more abundant and consistent members of the core gut microbiome of Crassostrea virginica when environmental oysters were compared with those depurated in sterile chambers for 5 days. (Pierce 2016). The abundance of Mycoplasma OTUs in bivalves has been documented in a number of studies (Green & Barnes 2010, Winters et al. 2011, King et al. 2012, Wegner et al. 2013, Cleary et al. 2015, Lokmer et al. 2016b, Arfken et al. 2017, Pierce & Ward in review). These obligate parasites are commonly found within a number of hosts, from humans to fish (Chen et al. 2012), so their presence in bivalves is not surprising, although their high abundance may be a sign of infection. Other common members of the core gut microbiome of C. virginica were Phaeobacter spp. and spirochetes in the family Brachy-spiraceae (Pierce 2016). Spirochetes have long been considered symbionts of the oyster gut because of their frequent isolation from the crystalline style (Margulis & Hinkle 1992). While Phaeobacter spp. have been implicated as critical for host defense against invading Vibrio pathogens (Prado et al. 2009, Karim et al. 2013, Sohn et al. 2016, Zhao et al. 2016), their naturally high abundance indicates a symbiosis with oysters. Despite the presence of potentially probiotic Rhodobacterales OTUs, vibrios are still present in oyster tissues and represent another candidate for the core microbiome (Vezzulli et al. 2018, Pierce 2016).

MICROBIAL ROLE IN BIVALVE PHYSIOLOGICAL ENANTIOSTASIS

Pathogen Susceptibility

Gut bacteria are of particular interest because of their ability to influence a variety of processes in their host, including nutrient absorption through enzyme production, immune system development, and disease susceptibility (Guarner & Malagelada 2003, Rawls et al. 2004, Forberg et al. 2012, De Schryver & Vadstein 2014, Dishaw et al. 2014a). As invertebrates, possessing only innate immunity, bivalves rely on humoral and cellular defense via a variety of molecules and processes including antimicrobial peptides, lysozymes, lectins, hemocytes (i.e., phagocytosis), apoptosis, and a respiratory burst resulting in the production of reactive oxygen species (Chu 1988, Canesi et al. 2002, Goedken et al. 2005, Pruzzo et al. 2005, Hughes et al. 2010). The microbial communities that animals harbor may also provide protection against pathogens, and changes in bacterial diversity and abundance have been linked to altered health statuses for a range of hosts (e.g., mice--Ley et al. 2005, Turnbaugh et al. 2006; corals--Frias-Lopez et al. 2002, Roder et al. 2014; shrimp--Rungrassamee et al.2016).

Because the gut is considered a major entry point for disease transmission, a diverse and established microbial community may protect the host from invading pathogens. Mechanisms of protection by the resident bacteria could include the secretion of antimicrobials, production of cell signaling molecules, the regulation of host genes, and the monopolization of resources (i.e., space and nutrients; also known as "colonization resistance") (Freter et al. 1983a,b, Cross 2002, Stecher & Hardt 2008, Kau et al. 2011, Forberg et al. 2012). Disturbance of the normal microbiota creates an opening for pathogens, in part, by making available formerly occupied niches (Mondot et al. 2013, De Schryver & Vadstein 2014). In humans, antibiotic use, and thus disturbance of the gut microbiome, has been linked to the development of intestinal infections by Clostridium difficile (Hall and O'Toole, 1935), inflammatory bowel disease, and allergies, to name a few (Levy 2001, Chang et al. 2008, Blaser 2011, see Ianiro et al. 2016 for review).

With respect to bivalves, a few studies have also made a connection between health and microbial diversity. Heat stress in Crassostrea gigas resulted in a decrease in hemolymph microbial diversity, followed by death after pathogen challenge (Lokmer & Wegner 2015). Sydney rock oysters, Saccostrea glomerata (Gould, 1850), infected with a protozoan parasite, Marteilia sydneyi Perkins and Wolf (1976), were found to have significantly reduced bacterial diversity in their gut compared with uninfected individuals (Green & Barnes 2010). It is unclear, however, if reduced bacterial diversity is caused by, or a result of, disease in many instances. Recently, the application of antibiotics was used to understand how reduced bacterial diversity of the gut influences Vibrio accumulation in eastern oysters (Crassostrea virginica). Whereas antibiotics, and thus gut microbiome disturbance, did not affect the overall levels of pathogen accumulation, it did significantly reduce the ability of oysters to reduce pathogen levels over time (Pierce 2016). This result suggests that a highly diverse community of gut micro-biota influences the ability of oysters to reduce pathogen loads. Meanwhile, no significant reduction in Vibrio loads in oysters from the low diversity treatment may be a result of the idea that pathogens were able to begin colonizing.

Ecologically, the diversity-invasion theory suggests that a diverse community may deter pathogen colonization, as has been seen in other pathogen-microbiome studies (Dillon et al. 2005). For example, altered microbial communities have been associated with pathogen progression in other invertebrate species. Disease challenges using Vibrio harveyi (Johnson & Shunk, 1936) on two species of shrimp resulted in the alteration of their normal intestinal microbiota (Rungrassamee et al. 2016). In addition, a difference in bacterial community structure was observed for corals infected with black band disease compared with corals without the disease (Frias-Lopez et al. 2002, Frias-Lopez et al. 2004).

Digestive Enzymes

Bacteria in the gut of organisms aid in the digestion of food and thus the absorption of nutrients by producing enzymes the host is not capable of producing, or by increasing the abundance of enzymes critical to digestion (see reviews: Guarner & Malagelada 2003, Flint et al. 2008). In the bivalve gut, enzymes that could be produced by bacteria have been observed, including amylases, cellulases, proteases, and lysozymes (Shivokene et al. 1986, Lawry 1987, Seiderer et al. 1987). Bivalves themselves may produce many of these enzymes, in addition to lipases and laminarases (Mathers 1973, Langdon & Newell 1996, Ibarrola et al. 1998, Arambalza et al. 2010, Adeyemi & Deaton 2012, Sauey et al. 2015), but most studies have not differentiated between the digestive enzymes produced by the host versus those produced by the microbiota. In Crassostrea virginica, however, a few studies have used antibiotics in an attempt to differentiate between host- and microbe-produced enzymes. Results of this research demonstrate that microbial communities have little or no effect on total enzyme production in the gut (Newell & Langdon 1986, Mayasich & Smucker 1987, Crosby et al. 1989). Although conflicting results showing that bacteria do in fact increase carbon assimilation in oysters have been reported (Crosby et al. 1990), the prevailing hypothesis is that oysters can produce cellulases and chitinases. In these studies, however, the effects of antibiotic treatment on resident bacteria were assessed using culture-dependent techniques. Thus, the actual impact on the microbial community of the oyster was unknown. In an attempt to determine the full effect of antibiotic treatment on the microbial community of the gut, Pierce (2016) exposed oysters to a trio of antibiotics for 3 days and then quantified changes in the microbiome by means of next-generation sequencing. Results indicated that application of the antibiotic cocktail reduced the number and diversity of bacterial taxa in oysters and the functional diversity (richness) of the microbiota. It also reduced the number of OTUs shared among oysters compared with controls. The alteration of the microbial community resulted in reduced production of the enzyme xylanase. It did not affect the production of cellulase, protease, or amylase enzymes, nor did it impact the absorption efficiency of organics from the diet. Furthermore, research on the pearl oyster Pinctada fucata (Gould, 1850) has shown the presence of genes coding for amylase and chitinase enzymes (Huang et al. 2016, Li et al. 2017), supporting endogenous enzyme production in bivalves.

Although the aforementioned studies have contributed important information, an understanding of whether the host, the resident microbial communities, or both produce critical digestive enzymes is nascent, and many questions remain. For example, one unknown is the contribution of enzymes from lysed cells, which has been shown to impact digestion even after bacteria are eradicated (Harris 1993). A second unknown is the contribution of bacterial-bacterial or bacterial-animal horizontal gene transfer. Several examples within sessile marine invertebrate phyla have emerged to suggest that the incorporation of bacterial genes into host animal genomes occurs and is related to improving metabolic function (Boto 2014, Degnan 2014). In this case, bacterial genes responsible for enzyme production and other metabolic processes could be transferred to the eukaryotic host genome. Finally, functional redundancy of enzyme production genes could occur within the microbial community. As such, the antibiotic elimination of certain bacteria may not result in an overall disruption of enzyme production.

As with disease, a disturbance in the gut microbiome has the potential to impact host enantiostasis with respect to digestive enzymes. A reduction in the host's nutrient absorption may result in a number of downstream consequences such as alteration of the host's energy budget and impact growth and reproduction. Further work is needed to elucidate conflicting results. Many unknowns could be probed more directly with the aid of complete genomes, which are largely lacking for bivalves.

FUTURE RESEARCH

The results summarized in this review provide a base of knowledge to further investigate microbial-host interactions in a meaningful way. Microbial communities make excellent models for testing ecological theory. Their rapid growth allows investigating questions in the short term that would take years or even decades to understand in eukaryotes (e.g., Lyons et al. 2010). Disturbance, succession, colonization, and diversity-invasion theories are readily testable by experiments using hosts and their associated microbiota (e.g., Dillon et al. 2005). One of the many challenges is comparing different studies with disparate results. Alterations in nucleic acid extraction, PCR, and amplicon-sequencing methods may account for differences among studies. In addition to methodological variations, the influence of host genetics and environmental conditions are factors affecting the microbiome, which are only beginning to be understood.

Regarding future studies on bivalves and their microbial communities, important next steps are to determine (1) how bivalve genotype and physiological status influence the microbiome, (2) how changes in the microbiome affect digestive function and other physiological processes, and (3) how changes in the microbiome affect immune status of the bivalve host. Population studies would help tease apart host and environmental influences on the microbial community. By detecting previously identified, oyster-specific single nucleotide polymorphisms, a profile of genetic variation of populations can be created (Quilang et al. 2007). Other work has shown a link between genetic diversity of the gill microbiota and genetic similarity of Pacific oyster hosts (Wegner et al. 2013), and analogous studies need to be conducted for a range of bivalve species. Long-term laboratory experiments to probe the bacterial-host dynamics of digestive enzyme functioning should be conducted to rule out the contribution of recently lysed cells. In addition, whole bivalve genomes would help to identify metabolic genes for endogenous enzyme production. Meta-genome sequencing and measuring immune functions (e.g., respiratory burst and phagocytosis) should be carried out to give an insight into how host response differs with antibiotic treatment. Genes related to the production of reactive oxygen species, lysozymes, and phagocytosis have been found in Crassostrea gigas and their expression has been linked to the presence of virulent vibrios (Fluery et al. 2009, de Lorgeril et al. 2011). If the resident microbiota influences the host immune response to infection, disturbing the community with antibiotics should result in altered immune-related gene expression in those oysters. Additional improvements to experimental design include measuring bivalve clearance rates and repeated challenges over time to compare acute versus chronic exposures.

As with most research, results only generate more questions. There are many more topics to examine relating to the bivalve microbiome. For example, do core microbiome trends hold true across bivalve species? Do they hold true with other suspension feeders (e.g., Crepidula spp.)? Are observed similarities between oyster and mussel microbiomes a result of their shared feeding mechanism? Are observed differences between oyster and mussel microbiomes a result of their genetic differences? Do other marine filter feeders harbor the same numbers and types of bacteria as bivalves? Do they share a core? With continued reference to the eastern oyster, do Crassostrea virginica from other locations (i.e., Chesapeake Bay and Gulf of Mexico) harbor the same bacteria as those in Long Island Sound? How far do spatial trends extend until they are broken? Does the environment impart more of an influence in some locations than in others? Do the core microbiota contribute to specific host physiological functions? Does a high diversity in the gut microbiome inhibit pathogen colonization in the long term? Investigating such questions would be beneficial, resulting in an enhanced understanding of bivalve host-bacterial interactions.

CONCLUSIONS

Understanding natural variation in the genetic and functional diversity of the oyster-associated microbial communities is vital for establishing a baseline to which the effects of extrinsic and intrinsic factors can be compared. The studies described here help to inform the growing fields of marine microbial ecology and microbiome research. Building on this work will allow more direct probing of questions relating to the role of microbial communities in host physiological functioning and enantiostasis. The complex relationships between the environment, bivalves, and maintenance of their microbial communities have only begun to be probed in the past 30 years. Many of the mechanisms that mediate prokaryote-host symbioses are unknown or unclear. Both extrinsic and intrinsic factors are at play, with no clear dominant influence. The potential for gut microbial communities to effect bivalve digestive enantiostasis and pathogen accumulation is great. Thus, understanding the natural spatial and temporal variation of these communities, the influence of the surrounding seawater and particulate-associated microbes, and the impact that disturbances in the microbiome have on bivalve enantiostasis is imperative. Because of the prominent role of bivalves in the environment, implications may exist not only on the individual and population level but also for ecosystem functioning. As bivalves are a key component in ecosystem health, the factors that influence oyster functioning and disease are critical to maintaining nearshore, benthic ecosystems. In light of changing climate regimes, it is crucial to understand the relationships between hosts and their associated microbiota, stressors, and environmental perturbations. By investigating specific host-microbiome interactions, ecological predictions for the future could be made.

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MELISSA L. PIERCE (1*) AND J. EVAN WARD (2)

(1) Department of Biological Sciences, University of Illinois at Chicago, Chicago, IL 60607; (2) Department of Marine Sciences, University of Connecticut, Groton, CT 06340

(*) Corresponding author. E-mail: mlp 16@uic.edu

DOI: 10.2983/035.037.0410
TABLE 1.
Summary table of alpha diversity metrics reported for studies on
bivalves using 16S next-generation sequencing to study host-associated
microbial communities.



Bivalve species            Location

Crassostrea virginica      Long Island Sound, CT

Crassostrea virginica      Atlantic Beach, NC
Crassostrea virginica      Gulf of Mexico, LA
Crassostrea virginica      Gulf of Mexico, LA
Crassostrea coteziensis    Gulf of California,
                           Mexico
Crassostrea sikamea        Gulf of California,
                           Mexico
Crassostrea gigas          Gulf of California,
                           Mexico
Crassostrea gigas          Wadden Sea, Germany
Crassostrea gigas          Wadden Sea, Germany
Crassostrea gigas          Wadden Sea, the
                           Netherlands and
                           Germany
Crassostrea gigas          Wadden Sea, the
                           Netherlands and
                           Germany
Crassostrea gigas          Wadden Sea, the
                           Netherlands and
                           Germany
Crassostrea gigas          Wadden Sea, the
                           Netherlands and
                           Germany
Crassostrea gigas          Wadden Sea, the
                           Netherlands and
                           Germany
Crassostrea gigas          Gulf of La Spezia, Italy
Crassostrea gigas          Gulf of La Spezia, Italy
Crassostrea hongkongensis  Hailing Bay, China
Mytilus edulis             Long Island Sound, CT

Mytilus edulis             Barnegat Bay, NJ
Mytilus edulis             Barnegat Bay, NJ
Mytilus galloprovincialis  Gulf of La Spezia, Italy
Mytilus galloprovincialis  Gulf of La Spezia, Italy
Brachidonles sp.           Kakaban and Maratua islands, Indonesia
Brachidontes sp.           Kakaban and Maratua
                           islands, Indonesia
Ruditapes philippinarum    Arcachon Bay, France
Ruditapes philippinarum    Arcachon Bay, France
Ruditapes philippinarum    Venice lagoon.

                           Water
                           temperature
Bivalve species            [degrees]C   Time of year

Crassostrea virginica       4-21

Crassostrea virginica      n/a
Crassostrea virginica      n a          August and September
Crassostrea virginica      n/a          August and September
Crassostrea coteziensis    26-29        July-September

Crassostrea sikamea        26-29        July-September

Crassostrea gigas          26-29        July-September

Crassostrea gigas           2
Crassostrea gigas           8 and 21    August and November
Crassostrea gigas          13-22        June-August


Crassostrea gigas          14


Crassostrea gigas          14


Crassostrea gigas          14


Crassostrea gigas          14


Crassostrea gigas          26.7
Crassostrea gigas          26.7
Crassostrea hongkongensis  n a         March-December, monthly
Mytilus edulis              4-21

Mytilus edulis             n a
Mytilus edulis             n/a
Mytilus galloprovincialis  26.7
Mytilus galloprovincialis  26.7
Brachidonles sp.           28-32
Brachidontes sp.           n/a          August

Ruditapes philippinarum     1-25        April and November
Ruditapes philippinarum     1-25        April and November
Ruditapes philippinarum     3.8-25      June, December, and January



Bivalve species                                  Tissue type

Crassostrea virginica      September, November,  Gut
                           March, and July
Crassostrea virginica      July                  Gut
Crassostrea virginica                            Gut
Crassostrea virginica                            Stomach
Crassostrea coteziensis                          Gut

Crassostrea sikamea                              Gut

Crassostrea gigas                                Gut

Crassostrea gigas          January               Gills
Crassostrea gigas                                Hemolymph
Crassostrea gigas                                Hemolymph


Crassostrea gigas          n/a                   Hemolymph


Crassostrea gigas          n a                   Gills


Crassostrea gigas          n/a                   Gut


Crassostrea gigas          n/a                   Mantle


Crassostrea gigas          August                Gut
Crassostrea gigas          August                Hemolymph
Crassostrea hongkongensis                        Gut
Mytilus edulis             September, November,  Gut
                           March, and July
Mytilus edulis             n/a                   Gut
Mytilus edulis             n/a                   Gills
Mytilus galloprovincialis  August                Gut
Mytilus galloprovincialis  August                Hemolymph
Brachidonles sp.           August                Whole organism
Brachidontes sp.           Whole organism        1,860

Ruditapes philippinarum    Gut                     142
Ruditapes philippinarum    Gills                   100
Ruditapes philippinarum    Gut                   2,170

                             Operational
                           taxonomic units  Shannon        Bacterial
Bivalve species            (OTUs) reported   index           phyla

Crassostrea virginica        781                4             11

Crassostrea virginica        477-552            1.06-1.33      2
Crassostrea virginica        243-304            3.96-4.05     12
Crassostrea virginica        138-172            1.27-3.63     12
Crassostrea coteziensis      117-368            3.2-4.5       13

Crassostrea sikamea           79-367            2.5-1.5       13

Crassostrea gigas            234-305            1.87-4        13

Crassostrea gigas          4,464                2.5-4         10
Crassostrea gigas          2,622                4.2-4.8       18
Crassostrea gigas            100            Avg 4.4          n/a
                                            and 3.8 at
                                                2 sites
Crassostrea gigas            500-600          n/a            n/a


Crassostrea gigas            200-400          n/a            n/a


Crassostrea gigas            100-300          n/a            n a


Crassostrea gigas            200-400          n/a            n/a


Crassostrea gigas            600              700 (*)        n/a
Crassostrea gigas          1,200            1,300 (*)        n/a
Crassostrea hongkongensis    n/a                2.2-2.8      n/a
Mytilus edulis               989              4-6.5           22

Mytilus edulis               178                1.4-4.0      n/a
Mytilus edulis                68                0.3-1.9      n/a
Mytilus galloprovincialis    600-700          800 (*)        n/a
Mytilus galloprovincialis  1,000            1,100 (*)        n/a
Brachidonles sp.           3,553              n/a             44
Brachidontes sp.             n/a               38             38

Ruditapes philippinarum      n/a              n/a            n/a
Ruditapes philippinarum      n/a              n/a            n a
Ruditapes philippinarum      500-1,100 (*)      7              7



Bivalve species            Reference

Crassostrea virginica      Pierce and Ward (in review)

Crassostrea virginica      Arfkenet al. (2017)
Crassostrea virginica      King et al. (2012)
Crassostrea virginica      King et al. (2012)
Crassostrea coteziensis    Trabal et al. (2014) ([dagger])

Crassostrea sikamea        Trabal et al. (2014) ([dagger])

Crassostrea gigas          Trabal et al. (2014) ([dagger])

Crassostrea gigas          Wegneret al. (2013)
Crassostrea gigas          Lokmer and Wegner (2015)
Crassostrea gigas          Lokmer et al. (2016a)


Crassostrea gigas          Lokmer et al. (2016b)


Crassostrea gigas          Lokmer et al. (2016b)


Crassostrea gigas          Lokmer et al. (2016b)


Crassostrea gigas          Lokmer et al. (2016b)


Crassostrea gigas          Vezzulli et al. (2018)
Crassostrea gigas          Vezzulli et al. (2018)
Crassostrea hongkongensis  Wang et al. (2016)
Mytilus edulis             Pierce and Ward (in review)

Mytilus edulis             Schill et al. (2017)
Mytilus edulis             Schill et al. (2017)
Mytilus galloprovincialis  Vezzulli et al. (2018)
Mytilus galloprovincialis  Vezzulli et al. (2018)
Brachidonles sp.           Cleary et al. (2015)
Brachidontes sp.           Cleary and Polonia (2018)

Ruditapes philippinarum    Meisterhans et al. (2016)[section]
Ruditapes philippinarum    Meisterhans et al. (2016)[section]
Ruditapes philippinarum    Milan et al. (2018)

n/a = not available.
 (*) Chaol index.
([dagger]) Data reported for adult oysters only.
([section]) Method utilized was ARISA, not sequencing.
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Author:Pierce, Melissa L.; Ward, J. Evan
Publication:Journal of Shellfish Research
Article Type:Report
Date:Oct 1, 2018
Words:13403
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