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. The immune system contributes to host and host-symbiont homeostasis by eliminating foreign particles such as viruses, infectious bacteria or parasites (Fournier et al., 2000). 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. The beneficial role of microbiota in bivalve species, and in particular the ability to protect against pathogens and environmental stressors, has recently been investigated (S Ossai et., al 2017; Lockmore and Wegner 2015). However, the role of resident bacteria in the host response to chemical contamination remains largely unexplored.
The Eastern oyster, 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). Pesticides are introduced into rivers by ground “scrubbing” when rainfalls occur and then may enter marine areas, particularly estuarine and coastal zones. These pollutants may have major ecological consequences and could endanger organism growth, reproduction or survival by fundamentally changing the associated core microbiota (Banerjee et al., 1996).
Here, 16s rRNA gene amplicon sequencing was used to characterize the microbiome of C. virginica. 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 μg/L levels. The presence of herbicides in aquatic environments is one of the major challenges for the preservation of this 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 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 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.
The U.S. Safe Drinking Water Act established the maximum contaminant level of Atrazine to be 3 µg/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, which may make it more susceptible to disease (Cragin et. al., 2011). 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 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 by running 16S sequencing in hatchery-reared spat. The objective of the present study was to analyse the microbiota of juvenile oyster specimens in order to test what the response of resident microbial communities is to enviornemntal atrazine introduction. Here, we aim at assessing the impact of long-term exposure to pollutants on microbial communities, as well as to evaluate the potential impact of changes of microbial communities on host xenobiotic evolution and susceptibility to environmental chemicals, two crucial but still unresolved questions in ecotoxicology.
Methods
Oyster Acquisition and Stabilization
250 oyster (Crassostrea virginica) spat were purchased from Horn-Point Laboratory in April 2016. 50 oysters, of similar size, shell width, and age were chosen from >1000 juvenile diploid oysters. The 250 chosen oysters were subsequently assigned into 4 color coded groups containing 50 oysters in each group (Green, Pink, Blue, Orange, and White). Each color group was then parted into 5 sub-groupings and were labeled as follows: “Group Color” - I, II, III, IV, or V” (ie. Green-I, Green-II, Pink-I, Pink-II, etc.). Each sub-group contained 10 diploid oyster juveniles. 3.0 mm square mesh sieves were used to separate each sub-group. No oyster was smaller than 5.0 mm long x 4.0 mm wide when placed in the mesh sieves. A large holding tank, which served as the in-lab microcosm, was filled with 300L of pressure filtered water at a salinity of 25 parts per thousand (ppt). The oysters were allowed to grow in the lab within this microcosm for a stabilization period of three months prior to atrazine exposure. Frequent water changes (25% twice weekly) were used in order to minimize a buildup of both ammonium and nitrate levels 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 three-month stabilization period, oysters were fed 6 L of a concentrated phytoplankton mixture of (Tetraselmis Chuii, isochrysis galbana, and Nannochloropsis oculata) approximately ~ 400,000 cells/mL) 3 times per week on Mondays, Wednesdays and Fridays. Before adding 6L of the plankton mixture, 6L of water from the oyster microcosm were removed.
Relevant tank water parameters were monitored and adjusted as needed by replacing old saltwater with new saltwater. The tank was consistently maintained to fit the water quality parameters outline in Table 1: