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\begin{document}

\title{The effects of atrazine on the microbiome of the Eastern
Oyster:~\emph{Crassostrea~Virginia}~ ~ ~ ~ ~ ~ ~}



\author[1]{Adrian Britt}%
\author[1]{kate_cav}%
\author[1]{Dr. Tara Scully}%
\author[1]{Benjamin McSweeney}%
\author[1]{Kavery Nivana Theethira Poonacha}%
\author[2]{bamcsweeney94}%
\author[2]{manuelaromero20}%
\author[2]{Abhilasha Achar}%
\affil[1]{George Washington University}%
\affil[2]{Affiliation not available}%


\vspace{-1em}



  
  \date{May 08, 2024}


\begingroup
\let\center\flushleft
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\selectlanguage{english}
\begin{abstract}
~Over the last few decades, it has become clear that microbiota are of
vital importance for the survival, homeostasis, and overall development
of marine mollusks, and that shifts in their microbiota can dictate
chances of survival into adulthood (McFall-Ngai et al., 2013). Rapid
improvements in the technology used to assess microbial communities have
led to an expansion of the breadth and scope of microbial ecology
research over the past 20 years. Here, 16S rRNA gene amplicons were used
to evaluate how microbial compositions shift in response to exposure of
environmentally relevant concentrations of atrazine found within the
Chesapeake Bay (30 \selectlanguage{greek}µ\selectlanguage{english}g/L, 20\selectlanguage{greek}µ\selectlanguage{english}g/L, 10\selectlanguage{greek}µ\selectlanguage{english}g/L and 3 \selectlanguage{greek}µ\selectlanguage{english}g/L respectively). The
Chesapeake Bay has suffered a dramatic loss of its native mollusk
species, the Eastern oyster (Crassostrea virginica). It is believed that
this loss is caused in part by contamination of the Bay with atrazine, a
herbicide that washes off of agricultural crops and penetrates into
surrounding surface and groundwaters following heavy periods of rain in
spring and summer months. Although, the manner in which atrazine hinders
the survival of oysters remains unknown, the present study investigates
how exposure of atrazine to C. virginica affects its core microbiome.
The dominant bacterial groups found within all groups included those
belonging to the order~\emph{Rickettsiales, Vibrionales,~}and
\emph{Alteromonadales,} family~\emph{Pseudoalteromonadacea,
Vibrionaceaea,~}and \emph{Planctomycetaceae}, and
genus~\emph{Pseudoalteromonas, Vibrio~}and~\emph{Nocardia}. Although our
results confirm that oysters maintain similar core microbial communities
over time, we also report a novel finding, namely, that oysters exposed
to atrazine concentrations as low as 3\selectlanguage{greek}µ\selectlanguage{english}g/L saw a significant loss
of~\emph{Vibrio~}and~\emph{Pseudoalteromonas}~species interactions and
underwent a subsequent colonization of Nocardia species. Because the
microbiome of C. virginica is vital for its homeostasis and survival,
this result suggests that the presence of atrazine in the Chesapeake Bay
is in part responsible for widespread decline in this species
population.~%
\end{abstract}%



\sloppy


\textbf{Introduction}

~~~~The largest estuary in the United States, the Chesapeake once teemed
with oysters. In fact, its name comes from the Algonquin word
Chesepioooc, which translates to the Great Shellfish Bay (3). The
National Oceanic and Atmospheric Administration (NOAA) now reports that
the native oyster population of the Chesapeake Bay has diminished to 1\%
the original value (6). The microbiotas of marine mollusks are vitally
important for their survival, homeostasis, and development (McFall-Ngai
et al., 2013). Widespread use of the herbicide atrazine has generated
much research into its toxicity in aquatic systems.~

~~~~Atrazine, which has been banned from use by the European Union is
nevertheless the second~most widely used herbicide in the United States,
with an estimated annual production of 76 million pounds. When applied,
atrazine acts as a chemical contaminant of both surface and ground
waters.~ Atrazine
(6-Chloro-n-ethyl-n'-(1-methylethyl)-triazine-2,4-diamine) is a
synthetic herbicide commonly used on crops like corn, sugar cane, and
evergreens, especially~during spring and summer months (Agency for Toxic
Substances and Disease Registry, 2003) and paired with significant
rainfall events, washes into the tributaries which carry it over oyster
reefs and eventually into the estuaries such as the Chesapeake Bay.~One
of the beneficial roles of microbiota in oysters is to protect against
pathogens and environmental stressors, evidence has recently been
investigated (S Ossai et., al 2017; Lockmore and Wegner 2015). However,
the response of resident bacteria in the host to chemical contamination
remains largely unexplored.

~~~~The Eastern oyster,~\emph{Crassostrea virginica}, is one of the most
frequently cultivated bivalve species in the world and is typically
reared in estuarine environments that have become increasingly
threatened by exposure to pollutants. Among pollutants,
herbicide/pesticide contamination of shellfish has become more common in
estuarine areas over the past several decades due, in part, to chemical
run-off from terrestrial agriculture (Banerjee et al. 1996, khan et al.
2017). The microbial communities of suspension-feeding bivalves include
both resident and transient microbiota. 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.

~~~~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 bac- terial concentrations in
bivalves are higher by an order of magnitude than those in seawater, and
dominance of culturable bacteria is classically linked to sea- sonality
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 com- munity (Romero et al. 2002, Winters et al. 2010, Wegner
et al. 2013, Trabal et al. 2014, Li \& Wang 2017).

Agricultural pollutants may have major ecological consequences and could
endanger organism growth, reproduction or survival by altering the
beneficially associated microbiota (Banerjee et al., 1996). In this
study, two separate 16s rRNA gene amplicon sequencing events were used
to characterize the microbiome of hatchery raised~\emph{C.
virginica~}juveniles. We sought to characterize how atrazine effects the
bacterial composition of juvenile oysters after having been exposed to
environmentally relevant concentrations of atrazine three times a week
for a period of one-month\emph{.~}Extending the analysis to resident
microbial communities offers a unique opportunity to understand how host
and resident bacteria altogether respond to chemical contaminations.
Countless xenobiotic compounds, including pesticides, pharmaceuticals,
and personal care products, among others, are continuously introduced
into the environment and have been detected at concentrations up to \selectlanguage{greek}μ\selectlanguage{english}g/L
levels in surface waters. The presence of herbicides in aquatic
environments is one of the major challenges for the preservation of a
bivalves essential microbial environment. ~One noticeable service that
microbiota provide for their hosts is protection from pathogens (Kamada
et al., 2013). Indeed, in compromised hosts or under unfavorable
environmental conditions, the symbionts themselves have been understood
to act as opportunistic pathogens (Garnier et al., 2007; Cerf; Bensussan
and Gaboriau-Routhiau, 2010; Olson et al., 2014). As disease prevelance
has a large impact on the population dynamics and evolution of affected
organisms (Altizer et al., 2003), it is important to understand how
environmental factors and their resulting environmental stressors affect
the composition and function of microbiota and the outcome of
host--microbe interactions in~\emph{C. Virginia}.

~~~~As suspension feeders, oysters interact significantly with living
and non-living particles in the seston, including bacteria, as they
filter large quantities of water per unit time. It is thus unsurprising
that they harbor an order of magnitude more bacteria than does the water
in which they live (Colwell \&Liston, 1960; Cavallo et al., 2009).
Next-generation sequencing, although by no means free of biases (Fierer
and Lennon, 2011; Sergeant et al., 2012; Cai et al., 2013) enables
detailed characterization of microbial community composition and
dynamics, including rare phylotypes (Huse et al., 2008) that can act as
a seed bank and mediate community response to environmental change
(Caporaso et al., 2012; Pedros-Alio, 2012; Sjostedt et al., 2012).~

~~~~~The U.S. Safe Drinking Water Act established the maximum
contaminant level of Atrazine to be 3 \selectlanguage{greek}µ\selectlanguage{english}g/L (EPA). However, a study
conducted by the USDA in 2006 found the concentration of Atrazine in the
Chesapeake Bay watershed to be 30 ug/L, 10x the maximum contaminant
safety level (USDA, 2006). While advertised as safe, independent studies
have shown atrazine to cause chemical castration in frogs (Hayes et.
al., 2010), and increased menstrual cycle irregularity in humans (Cragin
et. al., 2011). Results like these suggest that atrazine may also be
inducing changes in the bacterial composition residing within the
eastern oyster, an organism which regularly interacts with the substance
in its ecosystem. This constant exposure to pollution~may make the
keystone species more susceptible to disease (Cragin et. al., 2011).
Indeed, in other documented aquatic ecosystems, the effects of atrazine
have proven to be particularly pronounced. ~It has been documented that
exposures to concentrations as low as 0.1 parts per billion of atrazine
in surface water have adversely affected frogs in causing the male
gonads to produce eggs -- effectually turning males into hermaphrodites
(DeLorenzo et al. 2001; Lynn 2017). The effects of atrazine at more
environmentally realistic concentrations are far less clear, and the
potential uninterrupted and adjuvant effects resulting from use of
atrazine on the survival and growth of~\emph{Crassostrea virginica~}are
simply not known.~

~~~The Chesapeake Bay has witnessed staggering losses to oyster
populations over the past century, reported to be down by 97\% when
likened to early records (Chesapeake Bay Foundation 2016). Atrazine is
commonly used in and around agricultural fields in the Chesapeake Bay
watershed (USDA). For this reason, it was chosen to be the focus in this
study examining the effects of herbicide-induced bacterial composition
changes. The objective of the present study was to analyze the
microbiota of juvenile oyster specimens in order to test what the
response of resident microbial communities is to environmentally
relevant concentrations of atrazine. Here, we aim to assess the impact
of long-term exposure to pollutants on microbial communities, as well as
to evaluate the potential impact of changes in microbial communities on
host xenobiotic evolution and susceptibility to environmental chemicals,
two crucial but still unresolved questions in ecotoxicology.

\par\null

~\textbf{Methods}

~~~\emph{Oyster Acquisition and Stabilization}

Two~separate specimen acquisition stabilization events took place for
this study, one in the Spring of 2017 and one in the Fall of 2018. For
both events, 250 - 300 ~juvenile~\emph{Crassostrea virginica~}oysters of
similar size and weight were purchased from Horn-Point Laboratory. In
the first event, oysters were separated into five groups of 50 specimens
and randomly assigned to an exposure group (30 \selectlanguage{greek}µ\selectlanguage{english}g/L Atrazine, 10 \selectlanguage{greek}µ\selectlanguage{english}g/L
Atrazine, 3 \selectlanguage{greek}µ\selectlanguage{english}g/L Atrazine, 30 \selectlanguage{greek}µ\selectlanguage{english}g/L Acetone and a Control). For the
second event, a sixth exposure group was added: 20 \selectlanguage{greek}µ\selectlanguage{english}g/L Atrazine. 3.0 mm
square mesh sieves were used to separate each group. No oyster was
smaller than 5.0 mm long x 4.0 mm wide when placed in the mesh sieves.
For each stabilization event, a large holding tank was filled with 300L
of pressure filtered water and raised to a salinity of 25 parts per
thousand (ppt). For the first stabilization event, oysters were allowed
to grow in the lab within this stabilization tank for a period of three
months prior to atrazine exposure. For the second event, oyster groups
were only allowed a stabilization period of two weeks due to time
restrictions. Frequent water changes (25\% twice weekly) were used in
order to minimize buildup of both ammonium and nitrate within the closed
water system . In addition to frequent water changes, Kordon AmQuel Plus
Ammonia Detoxifier/ Conditioner and TLC Saltwater aquarium conditioner
were used in order to remove Nitrate, Nitrite and Ammonia as
needed.~~During the first stabilization period, oysters were fed 6 L of
a concentrated phytoplankton mixture of (\emph{Tetraselmis Chuii,
isochrysis galbana, and Nannochloropsis oculata}) approximately
\textasciitilde{} 400,000 cells/mL) every other day.~~During the second
stabilization period, oysters were fed 6 L of a concentrated
phytoplankton mixture of (\emph{Tetraselmis Chuii, isochrysis galbana,
and Nannochloropsis oculata}) approximately \textasciitilde{} 400,000
cells/mL) every day.~

~~~~Relevant tank~water parameters were monitored and adjusted as needed
by replacing old 25ppt saltwater with new 25ppt saltwater. The tank was
consistently maintained to fit the water quality parameters outlined in
Table 1:~~

~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~~\selectlanguage{english}
\begin{longtable}[]{@{}l@{}}
\toprule
\begin{minipage}[t]{0.97\columnwidth}\raggedright\strut
~

Tabel 1. ~ ~ ~ ~ ~ ~ Salinity (ppt) ~ ~ ~ ~Ammonium (ppb) ~ ~ ~ ~Nitrate
(ppb) ~ ~ ~ ~ Plankton Concentration

~ ~ ~ ~ ~

~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ 25 ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~~ ~ ~ ~
~ ~ ~ 0 ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~0 ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~ ~
\textgreater{} 9,000 cells/ml

~ ~ ~ ~\strut
\end{minipage}\tabularnewline
\bottomrule
\end{longtable}

~Water quality parameters were monitored using:

(1) {]}Salinity Refractometer - Salinity

(2) API Testing Kit - Ammonium levels

(3) API Testing Kit - Nitrate levels

(4) Mass Spectrophotometer at 654nm - Plankton Concentration~

(5) Hemocytometer count - Plankton Concentration

\emph{Assigning Experimental Groups}

~ ~ For both events, oysters were randomly separated into five groups
with 50 oysters in each. To prevent overcrowding of the organisms, each
of these groups was then divided into a sub-group of 10 oysters. 3.0 mm
square mesh sieves were used to hold each sub-group. Since the
administered doses of atrazine were diluted in acetone, the Acetone
30~\selectlanguage{greek}μ\selectlanguage{english}g/L control group was used in order to ascertain acetone was not
causing the observed effects within treatment groups. Each group of 50
oysters was assigned to one of six treatments: 0~\selectlanguage{greek}μ\selectlanguage{english}g/L Atrazine/Acetone,
3~\selectlanguage{greek}μ\selectlanguage{english}g/L Atrazine, 10~\selectlanguage{greek}μ\selectlanguage{english}g/L Atrazine, 20~\selectlanguage{greek}μ\selectlanguage{english}g/L Atrazine, 30~\selectlanguage{greek}μ\selectlanguage{english}g/L Atrazine,
and 30~\selectlanguage{greek}μ\selectlanguage{english}g/L Acetone.

\par\null

~~~{\emph{Exposures}}

{~~~~All oyster groups were kept in separate tanks to avoid any
potential cross contamination. In order to expose the oyster groups to
atrazine and acetone, each group was removed from its holding tank and
placed into a separate 4-liter glass tank containing 2 liters of newly
made ~seawater (salinty= 25 ppt). ~Atrazine was added to each 4-liter
glass tank according to the corresponding treatment concentration. In
order to mimic the~heavy rainfall pattern surrounding the Chesapeake Bay
(USGS: Chesapeake Region Climate/Precipitation Data), treated groups
spent a total of 3 hours submerged in the treated 2 liters~of saltwater
three times per week, every other day (Monday,~Wednesday, Friday).
Aeration~was provided in both holding and treatment tanks.~}

~~~~Following ~each stabilization period, the experimental groups were
introduced to their first exposure. Four individual 4-liter glass tanks
were used to hold different concentrations of a
atrazine/acetone-saltwater mixture ( 25ppt): 3\selectlanguage{greek}μ\selectlanguage{english}g/L atrazine, 10 \selectlanguage{greek}μ\selectlanguage{english}g/L
atrazine, 20 \selectlanguage{greek}μ\selectlanguage{english}g/L atrazine, 30 \selectlanguage{greek}μ\selectlanguage{english}g/L atrazine and, 30 \selectlanguage{greek}μ\selectlanguage{english}g/L acetone
~respectively. ~The treatment rounds were conducted for a total of
30-days. For one of the events, surviving oysters were allowed to
continue to grow for an additional 30-day recovery period.

~~~~In order to minimize any potential contamination of the each group's
respective holding tanks after the 3-hour exposure period, each group
was washed using a constant stream of pressure-filtered water for three
one-minute rinses. Oyster groups were then relocated to their respective
40-liter glass bio-cube tanks. The bio-cube tanks were set to have the
same parameters as the large holding tank.~

\emph{Examined Concentrations}

\emph{~~~~}This study focused on four varying concentrations of atrazine
commonly found within an oyster's environment. In order to reproduce
ecologically relevant concentrations of atrazine within our experiments,
data resources overseen by the EPA and USDA were examined and cross
referenced with pesticide and herbicide datasets taken by various
external sources (USDA 2006; Flood et al. 2015; Powell et al. 2017;
Hively et al. 2011; EPA 2017). Concentrations of atrazine were observed
to be highest in the upper tributaries of the Chesapeake Bay. The
following outlines the chosen concentrations and the reasoning for
choosing the concentration.~

\par\null

1)~30 \selectlanguage{greek}µ\selectlanguage{english}g/L Atrazine: The peak concentration of atrazine detected in any
sample within the Chesapeake Bay was 30 \selectlanguage{greek}µ\selectlanguage{english}g/L (station L0002488), which
is located in a tributary on the Eastern Shore shown in
\emph{Supplementary~Figure 1} \textbf{.}~(US EPA 2006)

2) 20 \selectlanguage{greek}µ\selectlanguage{english}g/L Atrazine: A half-way point in between~30 \selectlanguage{greek}µ\selectlanguage{english}g/L Atrazine and 10
\selectlanguage{greek}µ\selectlanguage{english}g/L Atrazine for data exploration purposes.

3)10 \selectlanguage{greek}µ\selectlanguage{english}g/L Atrazine: The general trend in the data indicates that the
main detections of atrazine are in the tributaries while significantly
lower concentrations have been found in the Bay itself. During June of
2011, atrazine concentrations from the upper Chop Tank River were
logged. The average of all recorded concentrations was 0.29 \selectlanguage{greek}µ\selectlanguage{english}g/L. The
highest recorded concentration was 10 \selectlanguage{greek}µ\selectlanguage{english}g/L (Chesapeake Bay Program;
USDA: Hively et al. 2011).

4) 3\selectlanguage{greek}µ\selectlanguage{english}g/L Atrazine: The EPA's Maximum Residue Limit (MRL) for atrazine is
relevant because it is the limit which the EPA has set forth regulating
our drinking water (U. S. EPA MRL).

5) 3\emph{0}\selectlanguage{greek}µ\selectlanguage{english}g/L Acetone: Atrazine stock solution was dissolved in a
100\% acetone solution. In order to assume continuity throughout the
experiment this treatment was added to assure that any noticed effect on
development was due solely to atrazine exposure.~

~~~ ~\textbf{~}

\emph{16S Sequencing}

~ ~ As stated previously, two separate sequencing events took place
during this study. The first was conducted on specimens from the first
stabilization and treatment periods. The second sequencing event was
conducted in specimens from the second stabilization and treatment
periods. The first sequencing event included five examined
concentrations: 30 \selectlanguage{greek}µ\selectlanguage{english}g/L Atrazine, 10 \selectlanguage{greek}µ\selectlanguage{english}g/L Atrazine, 3\selectlanguage{greek}µ\selectlanguage{english}g/L Atrazine,
3\emph{0}\selectlanguage{greek}µ\selectlanguage{english}g/L Acetone and a control and included a total of 16 samples
(six 3\emph{0}\selectlanguage{greek}µ\selectlanguage{english}g/L Acetone, three controls, three 3\emph{0}\selectlanguage{greek}µ\selectlanguage{english}g/L
atrazine, two 1\emph{0}\selectlanguage{greek}µ\selectlanguage{english}g/L and two 3\selectlanguage{greek}µ\selectlanguage{english}g/L. Specimens in each group were
sequenced both directly following the treatment periods and following a
30-day recovery period. The second sequencing event included six
examined concentrations: 30 \selectlanguage{greek}µ\selectlanguage{english}g/L Atrazine, 20 \selectlanguage{greek}µ\selectlanguage{english}g/L Atrazine, 10 \selectlanguage{greek}µ\selectlanguage{english}g/L
Atrazine, 3\selectlanguage{greek}µ\selectlanguage{english}g/L Atrazine, 3\emph{0}\selectlanguage{greek}µ\selectlanguage{english}g/L Acetone and a control and
included a total of 24samples (four per sample). ~ Amplicons were
performed on a paired-end Illumina HiSeq2500 platform to generate 250bp
paired-end raw reads, and then pretreated. Specific processing steps for
both sequencing events were completed as follows:~

1) ~Paired-end reads were assigned to a sample by their unique barcode,
and the barcode and primer sequence were then truncated. 

2) ~Paired-end reads were merged using FLASH
(V1.2.7,http://ccb.jhu.edu/software/FLASH/ ) ,a very fast and accurate
analysis tool to merge pairs of reads when the original DNA fragments
are shorter than twice of the reads length. The obtained splicing
sequences were called raw tags. 

3) ~Quality filtering was then performed on the raw tags under specific
filtering conditions of Trimmomatic v0.33
(http://www.usadellab.org/cms/?page=trimmomatic) quality control
process. After filtering, high-quality clean tags were obtained. 

4) The tags were compared with the reference database (Gold database,
\url{http://drive5.com/uchime/uchime\_download.html}) using UCHIME
algorithm (UCHIME Algorithm.
(\url{http://www.drive5.com/usearch/manual/uchime\_algo.html}) to detect
chimeric sequences, and then the chimeric sequences were removed. The
data output of the above steps is shown in Table 1.

\emph{Statistical~Analysis}

~~~~To characterize microbial communities and to perform functional
analyses, the following wrappers were
employed:~~summarize\_taxa\_through\_plots~(to produce the taxonomical
files and
charts),~alpha\_rarefaction~and~beta\_diversity\_through\_plots~(to
assess respectively the alpha- and beta-rarefaction diversity indices),
principal\_coordinates.py (to compare groups of samples based on
phylogenetic distance metrics).~To compare sampling treatments
(Control~versus all treatment concentrations) within each sampling
season, ANOVA analyses have been performed at~genus~level
(P-value~\textless{}~0.005; FDR~\textless{}~0.01) using Excel version
16.13.

\par\null

~\textbf{Results}

\par\null

\emph{First sequencing event:}

\par\null

\emph{Diversity of microbial communities and taxonomic richness}

~~~~Of the top ten most abundant~bacterial genera strains identified
from the first sequencing event, ~Vibrio, Clostridum, and
Pseudoalteromonas were detected at high levels, while Nocardia,
Fusobacterium, Mycobacterium and Planctomycetes were detected at lower
levels. The high incidence of Vibrio, Clostridium, and Pseudoalteromonas
species may indicate the influence of environmental factors in oyster
groups. Additionally, a variety of unknown, likely transient, genera
were also detected across all samples, henceforth these genera will be
referred to as ``others''.~\selectlanguage{english}
\begin{figure}[h!]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/Screen-Shot-2018-04-12-at-11-06-20-AM1/Picture2}
\caption{{Experimental samples are listed on on the x-axis. Each color represents
a bacterial genus and the length of the color blocks indicate the
proportion of the relative abundance of the taxonomy; Y-axis: Relative
abundance (\%). (R) indicates a specimen that rested post exposure for
30-days before DNA extraction and sequencing. (NR) indicates a specimen
that did not rest before DNA extraction and sequencing.
{\label{649745}}%
}}
\end{center}
\end{figure}

~~~~~\emph{Pseudoalteromonas and Fusobacterium} displayed a decrease in
relative abundance within all samples treated~with atrazine. A
subsequent increase in the relative abundance of\emph{~Nocardia}, a
genus known for containing oyster pathogens, coincided with the decrease
of~\emph{\emph{Pseudoalteromonas and Fusobacterium}}. Directly following
the 30-day treatment period,~\emph{Nocardia} was not detected within any
of the samples except for Atrazine3.5(NR), ~where it retained a minimal
percent of relative abundance (\textless{}1\%). However, after the
30-day rest period, N\emph{ocardia~}was found within the majority of the
atrazine treated samples, with a relative abundance percentages ranging
from 5-13\% (Fig 2). Conversely, acetone and control samples never
displayed an association with~\emph{Nocardia}~either before or after the
30-day rest period. The only samples where N\emph{ocardia}~was detected
were those treated with atrazine. All three concentrations gave rise
to~\emph{Nocardia~}colonization, with the highest relative abundance
seen in sample Atrazine30.2(R) (Fig 1).~Clostridium was found to be
highly reduced in terms of relative abundance within all atrazine
treated samples. After the 30-day recovery period clostridium dropped to
\textless{}2\% relative abundance in all sequenced experimental groups.~

~~~~Clostridium, Psuedoalternamons and Fusobacterium were found to be
highly reduced in terms of relative abundance within all atrazine
treated samples. Clostridium dropped to \textless{}2\% relative
abundance in all experimental groups both directly after the treatment
period and after the 30-day recovery period. Psudoalteromonas and
Fusobacterium were also found to undergo a large reduction in relative
abundance, However, the effects were more pronounced after the 30-day
recovery period (Fig 1). The observed trends of a reduced relative
abundance of Clostridium, Fusobacterium and Psudoaltermonas held true
for both of the control groups as well, but to a lesser degree.~

~~~~A Principal Coordinates Analysis (PCoA) (Fig. 3) performed using
unweighted UniFrac distances allowed to further analyse the factors
responsible for significant differences between groups of samples. Fig 4
shows a clear separation along the x-axis (PC1~= 23.40\% of variability)
between control samples (Ace\_, Con\_) from the remaining atrazine
treated samples. Conversely, along the y-axis (PC2~= 10.97\%) all post
30-day recovery period atrazine treated groups form a small cluster on
the upper left quadrant of the graph, indicating bacterial composition
similarities.~

~~~~Prokaryotic diversity varied most between oysters treated with 3
\selectlanguage{greek}μ\selectlanguage{english}g/L atrazine without rest and the non-exposed control groups (Fig. 2).
Notably,~\emph{Vibrio~}was found to be absent in all but one of the
atrazine treated groups (Atrazine10.2(NR)) (Fig. 1).
~\emph{Arcobacter~}was present in all non-exposed control groups, but
not present in the experimental groups exposed to only to acetone.~

~~~~\selectlanguage{english}
\begin{figure}[h!]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/Screen-Shot-2018-05-21-at-12-45-44-PM/Screen-Shot-2018-05-21-at-12-45-44-PM}
\caption{{Each point represents a sample, plotted by a principal component on the
X- axis and another principal component on the Y- axis, which was
colored by group. The percentage on each axis indicates the contribution
value to discrepancy among samples.
{\label{325903}}%
}}
\end{center}
\end{figure}

~~~~Finally, the taxonomic composition was cross referenced with a
phylogenetic tree in order to understand how bacterial groups compared
to each other in terms of composition relatedness. From Fig. 3, we see
that control and acetone samples are all more closely related to each
other than the rest of the samples are, indicating a core microbiome
present within the control and acetone oysters. Conversely, experimental
samples that were exposed to atrazine are more closely related to each
other, displaying far fewer relative abundance \% for the
genus~\emph{Pseudoalteromonas, Clostridium,
FUsobacterium~}and~\emph{Vibrio~}than the control samples (Fig 1).~\selectlanguage{english}
\begin{figure}[h!]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/Screen-Shot-2019-01-16-at-1-56-11-PM/Screen-Shot-2019-01-16-at-1-56-11-PM}
\caption{{rThis is a caption
{\label{238135}}%
}}
\end{center}
\end{figure}

\emph{Second Sequencing Event:}

\par\null\selectlanguage{english}
\begin{figure}[h!]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/Screen-Shot-2019-01-14-at-12-25-07-PM/top-10-genus}
\caption{{Abundance of bacterial genus' observed in each sample and across groups:
CD: Control, AD: Acetone
{\label{279061}}%
}}
\end{center}
\end{figure}

\emph{~~~~For the second sequencing~event, i}t is clear that the overall
abundance of the top 10 most abundant genera was significantly decreased
in specimens that were exposed to concentrations of atrazine higher than
10\selectlanguage{greek}µ\selectlanguage{english}g/L. The same decrease in abundance holds true when looking at the
bacteria on the species level. Bacteroides abundance dropped to
negligible levels in the 30\selectlanguage{greek}µ\selectlanguage{english}g/L of atrazine group's specimens.
Allobaculum, Oscillospira , Escherichia, Lactobacillus, and Akermansia
genus' all significantly dropped in a dose dependent like manner,
whereas Staphylococcus species remained relatively stable amongst
treatment groups.~

\par\null\par\null\selectlanguage{english}
\begin{figure}[h!]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/f--Bacteroidaceae-g--Bacteroides/f--Bacteroidaceae-g--Bacteroides}
\caption{{A significant difference was observed in overall abundance~of
Bacteroidaceae and Bactericides when compared to the control. Indeed, a
dosage dependent relationship can be observed, as specimens were exposed
to higher concentrations of atrazine, they responded by exhibiting less
abundance of both~Bacteroidaceae and Bactericides.
{\label{301052}}%
}}
\end{center}
\end{figure}

~\textbf{Discussion}

~~~~Results from studies employing molecular methodologies, including
NGS, have found that~Proteobacteria~and~Cyano- bacteria~are abundant
members of bivalve gut communities, regardless of host species (King et
al. 2012, Trabal et al. 2014).~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 threeCrassostrea~spp.
analyzed. In individual oysters,~Proteobacteriamay account
for~\textgreater{}50\% of the total abundance, with~Alpha~(a)-
and~Gamma~(g)-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~g~-Proteobacteria,Shewanella,~Pseudoaltermonas,~Oceanospiralles,
and~Vibrio~are major taxa reported in this study. Other dominant phyla
identified included those seen by a variety of studies examining
microbiomes in bivalves: 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).

\par\null

~~~~~These results indicate that oysters maintain similar microbial
communities over time, acting as a host for a specific set of core
microbiota. The study found that control groups maintained a common
community of microbes in their overall body system throughout the
experiemnt. Conversely, we observed significant compositional variation
between and within experimental groups in the
genus~\emph{Pseudoalteromonas},~\emph{Vibrio}, and~\emph{Nocardia}. The
presence of the genus~\emph{Nocardia}~is of particular interest as they
are known oyster pathogens which cause round yellow-to-green pustules up
to 1 cm in diameter to be displayed on the surface of the mantle, gill,
adductor muscle, and heart (Friedman 1998).~\emph{Nocardia}~were found
only within the atrazine treated groups. During the 30-day exposure
period no significant relative abundance was observed
for~\emph{Nocardia} in all three atrazine exposed treatment groups,
however, after the 30-day rest period a significant relative abundance
was detected. This suggests that atrazine selected for the subsequent
survival and colonization of~\emph{Nocardia}.~Conversely,
Psuedoalternamonas, Clostridium and fusobacterium species were found
in~lesser degree within atrazine treated samples when compared to the
controls.

\par\null

This finding indicates that atrazine exposure may be play a role in
facilitating an environment which does not allow for Psuedoalternamonas,
Clostridium and/or fusobacterium\emph{~}species\emph{~}to colonize
oysters effectively. ~

~ ~ Interestingly, populations of bacterial species which decreased in
the presence of atrazine continued to sustain low abundance \% after a
30-day rest period, indicating that atrazine may have lasting effects
(greater than 1 month) in the population dynamics of \emph{C.
virginica.~Bacterial species known to be pathogenic to~\emph{C.
virginia}~and thus related to immunity and defense were of particular
interest in order to uncover how atrazine is affecting the
molecular-bacterial basis of oyster tolerance/resistance to the
herbicide's toxic effects. As has been reported in other studies
(Westlund et. at., 2017; Romero et. al., 2002;~A Arfken et. al., 2017;
~Pierce 2016) our results confirm that oysters maintain similar
microbial communities over time. The health of~\emph{C. virginica~is}
impaired by inflowing storm-water runoff from nearby agricultural
fields, which carry large amounts of atrazine into estuarine waters,
however,~\emph{~t}he knowledge of oyster-associated bacterial community
response to biocidal runoff is scarce.}

\emph{}

The sequence data obtained could be used to compare oyster aquaculture
management strategies as well as aquaculture practiced in different
regions that may have similar or different climactic conditions. It is
also meant to facilitate a greater understanding of how atrazine, as a
persistant environmental condition effects oyster-prokayote
interactions. Further studies of this nature could reveal important
links between oyster farming, environmental factors, husbandry
strategies, as well as legal regulations currently governing the
surrounding areas of the Chesapeake Bay.~~

\par\null

Future work:

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 po- tential for gut microbial
communities to effect bivalve digestive enantiostasis and pathogen
accumulation is great. Thus, un- derstanding the natural spatial and
temporal variation of these communities, the influence of the
surrounding seawater and particulate-associated microbes, and the impact
that dis- turbances in the microbiome have on bivalve enantiostasis is
imperative. 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 envi- ronment, bivalves, and maintenance of their microbial
communities have only begun to be probed in the past 30 years. Although
this study and many others have contributed important information, an
understanding of whether the host, the resident microbial communities,
or both produce critical di- gestive 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 hori- zontal
gene transfer. Several examples within sessile marine invertebrate phyla
have emerged to suggest that the in- corporation 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 en- zyme 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 en- zyme production.

\par\null

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.

\par\null

\textbf{Conclusion}

~~~~Understanding natural variation in the genetic and func- tional
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 presence of atrazine in the Chesapeake Bay
may be selecting for pathogenic bacterial groups to reside within the
oysters of the Chesapeake Bay and its tributaries. The effects of this
compositional shift remain unclear, therefore, future research efforts
should focus on understanding the relationship between biocidal
herbicides and oyster-prokaryote interactions.~~Extended experiments
with more time points, as well as repeated challenges, would help to
further elucidate the role of oyster-associated microbial communities to
the overall physiological functioning of the host. This research
provides a vital baseline for future research aimed at understanding the
role gut microbes have in oyster physiology.~\(\)

\par\null

\(\)

\textbf{~~}\(\)\textbf{Acknowledgements}

We would like to thank Horn-point laboratory for providing the spat used
throughout the experiment. Dr. Tara Scully for acting as the lead
investigator for the project. The George Washington University, for
housing the project, and providing the funds necessary for it to be
carried out. And finally, to each of the authors for their hard work and
perseverance.~

\par\null

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~

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\textbf{}

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