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 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. 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 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 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 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, 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 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.
Methods
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 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 µg/L Atrazine, 10 µg/L Atrazine, 3 µg/L Atrazine, 30 µg/L Acetone and a Control). For the second event, a sixth exposure group was added: 20 µg/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 (Tetraselmis Chuii, isochrysis galbana, and Nannochloropsis oculata) approximately ~ 400,000 cells/mL) every other day. During the second 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) 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:
Tabel 1. Salinity (ppt) Ammonium (ppb) Nitrate (ppb) Plankton Concentration 25 0 0 > 9,000 cells/ml |
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