Effects of Atrazine Runoff on Chesapeake Bay Aquatic Life

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Effects of atrazine runoff on Chesapeake Bay aquatic life:

Risk assessment of atrazine on the blue crab

Caitlin Andrews

Russell F. Ford

David Lucero

Henrietta Oakley

Satish Serchan

Executive Summary

Atrazine is the most extensively used herbicide for control of weeds in agricultural crops in the United States (EPA, 2003), with an estimated annual production of 76 million pounds (Hayes et al., 2003). Atrazine is frequently detected in surface waters and has been known to affect reproduction of aquatic flora and fauna, changing the community structure as a whole (Stagnitti et al.,2001). Due to negative impacts on aquatic life, it has been banned in the European Union.

In the Chesapeake Bay area, declines in abundance and health in submerged aquatic vegetation have been linked to increase atrazine use around the Bay resulting in an overall decline in the fish and waterfowl productivity (Christopher et al., 1992). Blue crabs (Callinectes sapidus) are an ecological indicator for the Bay and play large roles in energy transfer from estuaries to the nutrient limited ocean environment. Blue crabs play an important role in controlling the trophic cascades within an estuary. Blue crab populations do not flourish in watersheds associated with agricultural land use and pollution (King et al.,2005). Atrazine reduces chlorophyll-a within primary producers, prohibiting photosynthesis and cell division in certain species of submerged aquatic vegetation and phytoplankton important for the survival of the blue crab.

Exposures to concentrations as low as 0.1 part per billion of atrazine in surface water adversely affects frogs by causing the male frog gonads to produce eggs – effectively turning males into hermaphrodites (Hayes et al.,2003). These effects have been shown in controlled laboratory studies as well as in the wild. Elevated rates of human prostate cancer have been shown in some studies of workplace exposure. Atrazine may act on amphibians by stimulating production of aromatase, an enzyme that is linked to the growth of cancers in humans.

This report finds reasons for concern over the possible impacts of atrazine runoff on the Chesapeake Bay’s flora and fauna. In addition, this report presents direct and indirect impacts of atrazine on the blue crab’s habitat and food sources. This report also highlights the adverse effects of atrazine on frogs and human health. The information can be used for developing management guidelines and legislation.

Problem statement:

Atrazine is a common non-point source pollutant in runoff entering the Chesapeake Bay, and is suspected of contributing to the decline of blue crab populations.

Background:

The safety of atrazine in the environment has been the subject of recent review by the Unisted States Environmental Protection Agency and legal action by the Natural Resource Defense Council. Numerous studies address the impact of atrazine on individual components of the Chesapeake Bay ecosystem, but none of which we are aware, addresses possible impacts at the system level. The Chesapeake Bay is already heavily impacted by the agricultural lands found within its watershed threatening the health of the ecosystem and the viability of its fisheries.

Goal/Purpose statement:

This report will discuss literature relevant to possible impacts of atrazine runoff on the Chesapeake Bay estuary, including sub-aquatic vegetation (SAV), phytoplankton, and fauna dependent on these primary producers, particularly blue crabs (Callinectes sapidus). The role of the chemical as an endocrine disrupter in amphibians and humans will also be assessed.

Objectives:

An investigation of the presence and sources of atrazine in the Chesapeake Bay watershed will be conducted through a review of the pertinent literature. A broad range of studies will be identified that exemplify deleterious non-target impacts of the herbicide. The current review by the US EPA on the safety of atrazine, will be the basis for the argument against the safety of atrazine, and the attempt to model the effects of the pesticide on aquatic ecosystems will link cause and effect to the blue crab. Evidence of endocrine disruption and abnormal sexual development in amphibians due to exposure to atrazine will be used to argue that there is larger threat than what has currently been identified. This will show that the direct effects of atrazine exposure has undergone little research, though this endocrine conversion pathway is shared with humans and other mammals, and may linked to increased cancer rates in humans.

The cascade of trophic effects caused by atrazine will be the main argument for the significant harm to the Chesapeake Bay marine health. An analysis of the blue crab life history, habitat, and food source; will link atrazine application in agricultural lands to reduced productivity in the blue crab population. It will show the negative effects atrazine has on blue crab habitat, development, food sources, and water quality.

Introduction:

Chesapeake Bay watershed

Figure 1: Spatial map of Chesapeake Bay Watershed (King et al.,2005).

The Chesapeake Bay watershed stretches across more than 64,000 square miles, is home to 136 million people (Burke et al., 2000), and encompasses parts of six states and the District of Columbia [Figure 1]. The Chesapeake Bay is historically an important source of fish and shellfish, with a large industry based on the blue crab.

Figure 2: Land use of 19 sub-estuaries of the Chesapeake Bay (King et al., 2005).

The Chesapeake Bay also has agriculture land widely distributed throughout its watershed, though most is found in Maryland (King et al., 2005) [Figure 2]. 27 million acres of farmland can be found in the small state, which has impacted the Bay by increasing nutrient runoff, erosion, and agricultural chemicals (among other things) that enter the watershed (Burke et al., 2000).

Atrazine:

Atrazine was first registered for use as an herbicide on December 1, 1958 (Steinberg et al., 1995), and is a pre- and post-emergent, broad-leaf herbicide that works by inhibiting the growth of the target weeds by interfering with the normal function of photosynthesis (Chapman and Stranger, 1992). Atrazine is used mainly to suppress weed growth in corn, sorgum, and sugarcane. Structurally, a molecule of atrazine [Fig.3] consists of chloro and amino functional groups that inhibit photosynthesis in broadleaf and grassy weeds.

The EPA estimates that corn production accounts for 86% of the domestic usage of Atrazine. Approximately 75% of the field corn acreage in the U.S. is treated with Atrazine. Methods of application include groundboom sprayer, aircraft, and tractor-drawn spreader.

Atrazine in soil environment

Atrazine binds well in a soil profile after initial application. Clay and loamy soils have more affinity to atrazine than sandy soil because of soil aggregate structures and particle size density. Soil binding of atrazine initiates degradation of atrazine in the soil environment. The degradation of atrazine into derivative metabolites depends on factors including the soil type, percent organic matter, clay content, soil pH, and soil structure (Stagnitti et al., 1998; Kookana et al., 1998). There are five processes that determine the rate of atrazine degradation: hydrolysis, adsorption, photodegradation, volatilization, and microbial degradation (Stagnitti et al., 2001). Each of the degraded metabolites has varying degrees of persistence and toxicity (Stagnitti et al., 2001).

Fate of atrazine in aquifers and soils has been well documented in the past and many researchers have indicated that significant amount of atrazine may be stored in soil after application (Kookana et al., 1998). Atrazine is moderately hydrophilic, and a significant proportion is found in groundwater and surface runoffs due to its solubility and leaching potential (Stagnitti et al., 1998). Some research has shown the persistence of degraded atrazine metabolites in aquifers for more than 20 years (Hayes et al.,2003). Recent studies on the fate of atrazine in surface waters indicate the widespread occurrence of atrazine year round (Dana et al., 1993). One study found the concentration of atrazine as high as 49,000μg/kg in eroded soil leading to concentrations in surface waters as high as 1000 μg/L (Douglas et al., 1993).

Atrazine usage around the Chesapeake Bay area has increased exponentially and the concentrations of Atrazine in surface waters of the Chesapeake watershed can reach up to 98 μg/L in one growing season (Hall et al.,1999). In some agriculturally dominated regions concentrations can exceed 100 μg/L and persist for at least 30 days (U. S. EPA 2007).

Approach:

Research was conducted through the GoogleScholar literature service, using keywords “atrazine”, “Chesapeake Bay”, “blue crab”, “aquatic impacts”, “EPA”, “NRDC”, and using all two and three-word keyword combinations. ScienceDirect was also used, with the key words “atrazine,” “Chesapeake Bay,” “agriculture,” “blue crab,” “herbicide,” and “hypoxia.” ISI Web of Science was also used with keywords “blue crabs,” “mesozooplankton,” and “herbicide.” The U.S EPA website was searched using similar terminology, and considerable time was spent in tracking back the body of recent EPA decisions on atrazine regulation to the primary literature cited in those reports.

Findings:

Blue crab: Introduction

Blue crabs are important indicators of the Chesapeake Bay’s ecological health, and being both predator and prey, they serve important functions in the trophic cascades. Blue crabs are known for their complex migratory life cycle [Figure 4] and they exist over a wide range of habitats within the Chesapeake Bay (King et al., 2005).

Figure 4: Complex Migratory Lifecycle of the blue crab (Hines et al.,2008)

Habitat shifts are dependent on a variety of factors including salinity, food source, habitat, and location of mates (Hines et al.,2008; Dittel et al.,2008). Due to these migrations, blue crabs serve as important source of energy transfer in aquatic ecosystems: they take in energy and nutrients as a top predator within the Chesapeake’s estuary, and act as important food source for larger benthic mammals in the nutrient depleted pelagic ocean (Aguilar et al., 2008, King et al., 2005). Blue crabs also serve as good indicators of ecosystem health because of their traceable changes in spatial distribution due to anthropogenic effects on food and habitat (King et al.,2005; Seitz et al., 2005). Blue crab populations do not flourish in watersheds associated with agricultural land use. Watershed land use has been linked to reductions in fish community biodiversity (Moerke & Lamberti, 2006, King et al., 2005). Shore line alterations that eliminate the habitat of the blue crab, including marshes and downed woody debris, change the spatial distribution of the species (Seitz et al., 2005).

Due to blue crabs importance as ecological indicators in the Chesapeake Bay habitat, it is important to understand how atrazine affects the vitality of this species. The effects of atrazine on blue crabs are indirect, and thus it is important to have a thorough understanding of blue crab’s life cycle at all stages, including habitat and food sources.

Blue crab: Life Cycle

Blue crabs (Callinectes sapidus) range from Nova Scotia to Northern Argentina. Within the United States blue crabs are most common south of Cape Cod and mate along the estuaries of the mid-Atlantic seaboard, most abundantly in the Delaware and Chesapeake Bay (Aguilar et al.,2008, Davis & Davis, 2008). As with all species of crabs (Brachyura), blue crabs must molt as their internal tissue and body size increases, several times throughout their life (AlaskaFishScienceCenter, 2008). Callinectes sapidus females molt 18 to 20 times and males molt 21 to 23 times throughout their life time, not counting larval molts (Zinski, 2006).

The life cycle of blue crabs can be broken into three stages: larval, juveniles, and adults [Figure 5]. Female adults undergo a complex migration to mate and spawn (Hines et al.,2008 & Aguilar et al., 2008). Mating occurs from May to October in the low-salinity oligohaline zones of the upper estuary where males live. Migration to coalesced mating zones is often necessary (Hines et al., 2008). After mating, males will remain in these low-salinity zones while females may migrate in excess of 200 km to the mouth of the bay to zones of high salinity that are necessary for the first stage of larvae. Peak spawning occurs from May to August (Aguilar et al., 2008). Maximum lifespan for blue crabs is three years.

Figure 5: Patterns of migration of Blue Crabs (Hines et al.,2008)

The larval stage can be broken into zoeae and megalopae phases. The duration of the larval period ranges from four to six weeks (Tilburg et al.,2007). Spawning and release of larvae occurs primarily from July to August (Aguilar et al.,2008). Zoeae measure approximately 0.25 mm to 1.0mm, are filter feeders, and live in waters that are high in salinity (Zinski, 2008 & Tilburg et al.,2007). Areas of high salinity are found at the bottom of the estuary, near the mouth of the bay, and within the Atlantic Ocean. Blue crab larvae are found in large quantities in the Mid Atlantic Bight (MAB), a stretch of the Atlantic coastal ocean whose unique currents are influenced by salinity and buoyancy changes from the Chesapeake and the Delaware estuaries (Tilburg et al.,2007). Unlike many larvae, Callinectes sapidus do not exhibit vertical migration in the water column to counteract the ebb and flow of the ocean that could drag them out to sea. Rather the larvae remain near the surface among plankton throughout their zoeal development (Epifanio et al.,1989). Once the larval stage is complete, blue crab juveniles seek refugia within the estuary (Dittel et al.,2008).

Currents of the MAB exhibit a circulatory pattern in the summer months, transporting larvae southward close to shore, and northward, due to winds, further out to sea (Epifanio et al.,1989; Jones & Epifanio, 1995). Eventual transport into the natal estuary is controlled by downwelling and other wind-driven events that occur during autumn (Jones & Epifanio, 1995). Sanctuaries of null flow also exist with the MAB that prevent larvae from being transported long distances from the estuary (Tilburg et al.,2007). By the time Callinectes sapidus reenters the estuary it has reached its megalopae phase (Dittel et al.,2006).

Habitat

Blue crabs exist over wide range of habitats through their different life stages. Susceptible juveniles primarily rely on seagrass dominated-submerged aquatic vegetation communities for refugia and as a valuable habitat for their prey. Loss of seagrass beds as habitat for blue crab juveniles may force them upstream to lower-salinity waters with fewer predators and fewer food sources (Posey et al., 2005). Adult blue crabs continue to live in the salt marsh and marsh creek environment in order to survive, feed, and mate (Ryer et al.,1997). Damage to these seagrass dominated–submerged aquatic vegetation by atrazine is potentially devastating to blue crab populations.

Submerged aquatic vegetation (SAV) is an assembly of rooted macrophytes, dominated by seagrasses, found in the headwater of Chesapeake Bay’s tributaries. SAVs contribute to high primary and secondary productivity. Zooplankton feed on decaying grasses, barnacles, sponges, and amphipods. Therefore, SAVs serve as valuable source of refugia for juvenile fish and crustaceans, including blue crabs. These plant species also absorb excess nutrients and sediments, prevent shore erosion, and oxygenate the water (Kemp et al., 1984).

Blue crabs enter the Chesapeake Bay’s seagrass based estuaries during the megalopae phase. Megalopaes and young juveniles gradually migrate into the less-saline waters of the upper estuary. Megalopaes are susceptible to strong wind and water currents that might bring them further upstream (Pardieck et al.,1999). Settlement occurs in late summer and autumn. The juveniles will over-winter in these nursery habitats where they are protected from predators and food resources are abundant (Dittel et al., 2006). Megalopae are responsible for selecting a secure habitat, which will influence their survival into adulthood (Montfrans et al.,2003; Orth et al.,2002; Hovel et al.,2005). Megalopae and juveniles have been known to occupy sand, marsh mud, live oyster beds, and different types of seagrass communities including eelgrass (Zostera marina), smooth cordgrass (Spartina alternifolia), widgeongrass (Ruppia maritime), and shoalgrass (Halodule wrightii) (van Montfrans et al.,2003; Hovel et al., 2005; Orth et al., 2002; Moksnes et al.,2006).

Seagrass habitats, especially, that of Zostera marina, show the highest occupation of megalopaes and juveniles. Many juveniles will leave the sea grass community during the night, but a large majority of blue crab youngchoose this environment to overnight in (van Montfrans et al.,2003). Juveniles also show a preference for widgeon grass communities (Pardieck et al.,1999). High shoot density eelgrass communities provide the highest survivorship amongst blue crabs juveniles when compared to other sea grass communities (Orth et al.,2002). As a standard young blue crabs seek habitats with structural complexity both at local and landscape level scales. Three dimensional sea grass communities are uniformly preferred over mud habitats (Moksnes et al.,2006), and patchy sea grass communities are preferred over continuous sea grass environments (Hovel et al.,2005).

Maturity is reached after 20 post larval molts, around an age of one and a half years. Females cease to molt once they reach sexual maturity, while males have the ability to molt indefinitely (Zinski, 2008). Adults who continue to molt are much more susceptible to predation while their shells are growing. Molting adults rely on habitats such as seagrass marshes and the edges of marsh creeks to evade predation, especially during low tides (Ryer et al.,1997).

Submerged aquatic vegetation important to blue crab as habitat and refugia has shown sensitivity to atrazine in controlled experiments. Eelgrass is sensitive to full plant exposure of atrazine. Exposure of atrazine in groundwater to the root-rhizome has little effect on the species (Schwarzschild et al., 1994). If atrazine is present in the surface water there is a potential for detrimental effects based on length and amount of exposure. Acute exposure to atrazine (6 hours) at both 10μg/L and 100μg/L slowed the metabolic state of the plant. Net productivity decreased at an exposure of 100 μg/L. Chronic exposure of 100μg/L for twenty one days resulted in 50% mortality of the species. At lower levels, <10μg/L, chemicals that controlled the metabolic activity within the species actually increased their numbers in what is assumed at an attempt for survival (Delistraty et al., 1984). Widgeon grass communities experienced a 1% decline in photosynthesis at an exposure of 20μg/L and a photosynthetic reduction of 50% at an exposure of 95μg/L (Johnson et al., 1995). Smooth cord grass (Spartina alterniflora) exposed to atrazine for 35 days, showed little aversion to atrazine at amount of 3.1 mg/L (Lytle et al., 1998). These seagrass communities are important to blue crab development, and the sensitivity of these plant communities to atrazine poses a threat to their survival.