Effects of atrazine runoff on Chesapeake Bay aquatic life: Risk assessment of atrazine on the blue crab

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 young choose 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.
What Blue Crabs Eat

It is during the megalopae and juvenile phase the Callinectes sapidus changes its diet from a passive filter feed to actively seeking a more diverse omnivorous diet. The species continues to feed on phytoplankton and submerged aquatic vegetation (Dittel et al., 2006). Newly molted juveniles tend to feed on phytoplankton and zooplankton. Isolated juveniles crabs that were only fed zooplankton showed the fastest growth rate when compared to a number of other diets. Phytoplanktonic food webs also tend to dominate habitats that are within close proximity to the main estuary of Chesapeake Bay. Blue juvenile crabs feed on a variety of sources including, but not excluded to, submerged aquatic vegetation, marsh detritus, benthic algae, phytoplankton, amphipods, fiddler crabs (Uca pugnax), marsh periwinkle (Littoraria irroata), and thin shelled clams (Dittel et al., 2000; Dittel et al., 2006). High-density clam populations found in up-river sand and mud flats supported the most juvenile blue crab growth (King et al., 2005; Seitz et al., 2005).

Blue crab trophic position does not change with increasing body size and all size classes appear to consume primary producers and primary consumers in equal proportions (Hoeinghaus et al., 2007). Older juvenile crabs and adults are more likely to feed on sea grass detritus and benthic algae derived food webs. At the base of these food webs are C4 photosynthesizing plants, such as Spartina alternifolia, which are fed upon by omnivores such as marsh periwinkle and fiddler crabs. These food webs are most common in marsh zones, further from the main estuary (Dittel et al., 2000; Hoeinghaus et al., 2007). Blue crabs are also known to consume amphipods off macroalgae beds in the estuary. These amphipods graze directly on the macroalgae (Dittel et al., 2000). While adults continue to be omnivorous, they show preference for primary consumers including numerous bivalve species including soft clam (Mya arenaria) Atlantic rangia (Rangia cuneata) hooked mussel (Ischadium recurvum), and Balthic clam (Macoma blathica) (Kuhlmann et al., 2005; Ebersole et al., 2005).
Atrazine affects what blue crabs eat

While in its juvenile stage the effects of atrazine have the highest potential to negatively affect blue crab’s diet as majority of a larval blue crab’s first few weeks are spent in the coastal ocean, where atrazine concentrations are low and do not adversely impact the number of phytoplankton and zooplankton that blue crab zoeae feed upon. Megalopae and juveniles live in estuaries in and close to the mouth of the nine large rivers and numerous other creeks that flow into the Chesapeake Bay, where atrazine concentrations are highest (Pardieck et al., 1999; USGS 2004).

Phytoplankton forms the base of a food chain within the Chesapeake Bay estuary. 354 species of phytoplankton have been identified in the estuary, including 175 species of diatoms, 35 species of dinoflagellates, 78 chlorophytes, 35 cyanobacteria, nine species of euglenoids, and three species of prasinophytes. Diatoms, chlorophytes, and cyanobacteria are most abundant in the tidal fresh water of the estuary, while a different assemblage of diatoms, plus dinoflagellates and cryptomonads become more dominant in the lower estuary and saltier waters (Marshall et al., 1996). The bloom and bust cycles of these species of phytoplankton are complicated with a few species dominating the resources in different sections of the estuary. As a whole, the estuary experiences phytoplanktonic blooms in the summer and fall, with smaller bloom episodes in the spring. Freshwater and oligohaline species experience a higher cell maximum than waters downstream. The diatom (Skeletonema potamos) blooms in early spring, cyanobacteria (Chroo-coccus limneticus), (Merismopedia tenussimi), and diatoms C. striata, Stephanodiscus sp., Aulacoseira distans dominate in the summer, and the fall pulse consists of diatoms Asterionella formosa and S. potamos as the dominant species. Downstream cycles are dominated by diatoms Skeleonema costatum, Asterionella glacialis in the spring, summer and fall, and dinoflagellates Heterocapsa triquetra, Katodinium rotundatum, and Prorocentrum minimum, as well as several cryptomonads in the summer and fall (Marshall et al., 1996; Marshall et al., 1993).

Numerous studies have been performed to analyze the effects of atrazine on primary producers, as well as the effects of atrazine on riverine and estuarine ecosystems. Algae are perhaps the most susceptible aquatic organisms to atrazine (Solomon et al., 1996). At the molecular level numerous species show adverse effects to the exposure of atrazine. Chlorophyll content of (Chlorella pyrenoidosa) a freshwater chlorophyte, was greatly reduced, and cell division was prohibited when exposed to atrazine concentrations of 25μg/L and 50μg/L (Gonzalez-Murua et al., 1985). Reduction of chlorophyll-a affects photosynthetic intake and the ability of the organism to synthesis necessary nutrients (DeLorenzo et al., 1999). Growth inhibition was witnessed for numerous species of freshwater algae at exposure levels as 10μg/L in a closed controlled study. Green algae species Chlamydomonas sp., Chlorella sp., Pediastrum sp. and Scenedesmus quadricauda experienced evident growth inhibition and a reduction in chlorophyll at 100μg/L. Diatoms (Cyclotella hamma, C. meneghiniana, Synedra acus, and S. radians experience similar symptoms, but at a higher exposure (250μg/L) to the chemical. Some species, in particular Chlamydomonas sp. and Synedra acus experienced growth when exposed to low doses of atrazine (<10 μg/L) and an increase chlorophyll a activity (Tang et al., 1997).

The growth inhibition induced by atrazine on certain algal species has been proven in controlled experiments in estuarine mesocosms exposed to atrazine concentrations from 40μg/L to 160μg/. While many taxa of phytoplankton suffered from exposure to atrazine, especially large dinoflagellates, cyanobacteria species flourished, and large ciliates and small flagellates increased after 48 hours of exposure and remain elevated (DeLorenzo et al., 1999). Species of green algae almost uniformly have a lower threshold for atrazine than species of diatoms. This could have serious impacts on community structure and seasonal successional patterns of phytoplankton (Tang et al., 1997). Some have argued that exposure to atrazine does not reduce total bacteria and algal activity (Pollehne et al., 1999). Quick recovery of phytoplankton from an exposure ranging between 5μg/L to 20μg/L has been widely proven (Stevenson et al., 1982). Even populations of phytoplankton that were exposed to 1000μg/L of atrazine did not exceed an 80% loss, allowing for eventual for recovery (Tang et al., 1997). While recovery of the phytoplanktonic community is possible, the structure, and therefore function of the phytoplanktonic community does change when exposed to atrazine at least as long as exposure to the chemical lasts, plus recovery time. This change in structure may impact higher trophic levels (DeLorenzo et al., 1999).
Zooplankton and other fauna

Calanoid copepods (Acartia tonsa) which are abundant during the summer months are a prominent trophic imtermediary between phytoplankton to blue crabs. Calanoid copepods are filter feeders that thrive on dense plankton environment. As their density is directly correlated with the abundance and presence of phytoplankton, studies have shown reduction in reproduction and growth of these copepods as atrazine directly or indirectly affects their phytoplankton food source. A study on acute and chronic toxicity of atrazine on saltwater zooplanktons indicated that Acartia tonsa was the most sensitive species with an (acute toxicity) 96-h LC50 of 0.094 mg/L (G. S. Ward et al., 1985). Similarly, water flea Daphnia magna showed reduction in survival rate at 15 ppb (µg/L) and at 500 ppb the density of water flea in their natural environment decreased significantly (Stagnitti et al., 2001). As these abundant species decrease in numbers other opportunist zooplanktons get established resulting in change community structure and composition thus affecting blue crab community

.

Hypoxia

Coastal and estuarine hypoxia are impacting aquatic ecosystems world-wide, and the Chesapeake Bay is no exception. Seasonal oxygen depletion has been a problem for years, and nitrogen compounds are presumed to amplify this by limiting the photosynthesis of aquatic organisms (Adelson et al., 2001). The high agricultural land use has a negative correlation with coastal marsh and coastal seagrass habitats; agriculture is commonly associated with eutrophia and hypoxia (King et al., 2005), as fertilizers; pesticides and herbicides, provide a load of nitrogen and phosphorus to the waterways. A concentration dependent reduction in oxygen production can be seen in Elodea canadensis when exposed to atrazine (Vervliet-Scheebaum et al., 2007), and oxygen production in species of algae like Pseudokirchneriella subcapitata is also sensitive to the herbicide (Yeh and Chen, 2006). Atrazine has a direct effect on the production of oxygen of many aquatic primary producers, lowering the oxygen production in an already limited environment.

Figure 6. Episodic and chronic hypoxia; images for July 27 and August 3 represent chronic hypoxic conditions, while July 31 and August 1 represent a periodic upwelling event (Eggleston et al., 2005).

Much of the current concerns about the effects of atrazine stem from the research of Dr Tyrone Hayes. Atrazine had long been believed to cause harm to animals only at artificially high doses. During the 1990s while working for Syngenta, the largest manufacturer of atrazine, Hayes attempted to study the effects of atrazine at the low doses experienced in the aquatic environment in agricultural areas. At a level of .1 ppb, one-thirtieth of the EPA’s 3 ppb maximum contamination level for drinking water, the frogs showed sexual abnormalities, developing both male and female sex organs (Hayes 2002a). Atrazine is commonly found at these concentrations in drinking water and public waterways (van Dijk, 1999). In fact, even rainwater in the agricultural Midwest may contain atrazine at these levels (Hayes 2002a).

Hayes provided his report to Syngenta, which discounted his concerns and asked him to repeat the study. They denied his request to forward his results to the EPA. Hayes severed his relationship with Syngenta and repeated his frog study independently, as published in the Proceedings of the National Academy of Science (Hayes 2002a). Hayes’ work is of particular concern for two reasons. First, given the pervasiveness of the chemical in the environment, often at levels even higher than he used in his tests (Battaglin et al., 2000) it raises concerns for the health of amphibians, and even humans. The abnormalities he found were not externally visible, and had not been searched for in frogs or other taxa in the wild, although amphibian population declines generally are well recognized and still unexplained. Second, Hayes posits a mechanism for the sexual deformities in frogs involving suppression of testosterone and induction of estrogen production; the same endocrine conversion pathway involving the enzyme aromatase is present in humans and other mammals. In humans, aromatase has been shown to play a role in the formation and growth of cancers in breast and uterine tissue (Jongen et al., 2005).

Hayes and others have documented the same abnormalities in the wild, in leopard frogs (Rana pipiens) living near atrazine sources (Hayes, 2002b). Abnormal aromatase activities and functions have been shown from similar low exposures in salamanders, turtles, and fish (cited in NRDC 2007). Hayes’ research came in the middle of an Environmental Protection Agency evaluation of atrazine, and caused a welter of accusations about the validity of his work. Syngenta hired a new team to replace Hayes and repeat his frog studies. These later studies do not appear to have been peer-reviewed or published, and access to them was not permitted for this paper. EPA (2003b pp 6-7) cites 17 studies examining atrazine disruption of amphibian sexual development, the majority industry-supplied and unpublished.

The EPA has reached several decisions. First, it states that even if atrazine is shown to be an endocrine disrupter in the environment at actually occurring levels, this does not constitute an ‘endpoint’ requiring action. Second, the EPA states “None of the studies fully accounted for the environmental and animal husbandry factors capable of influencing endpoints that the studies were attempting to measure” (EPA, 2003a,b). Finally the EPA concludes that there was not enough consistent evidence to confirm that atrazine alters amphibian development (EPA, 2003 a,b).
Threats to Human Health

Equally contentious is the attempt to evaluate human cancer risks from epidemiological study of workers exposed in the manufacture and use of atrazine. Detailed studies by several parties have been made of the incidence of prostate cancer and of Non-Hodgkin’s Lymphoma among employees of a St. Gabriel (Louisiana) chemical plant that produces atrazine. Assertions in this case center on attempts to discern whether a higher-than-expected rate of prostate cancer among employees is due to workplace exposure, or whether it is due to the success of a more rigorous screening process (EPA 2003 b, pp 3-6). The EPA conclusions align with those made by Syngenta, that there is no significant statistical correlation between exposure and prostate or other cancers.

The Natural Resource Defense Council (NRDC) disagrees, however, and counters with a 28 page paper titled “Atrazine: An Unacceptable Risk to America’s Children and the Environment” (attachment B in EPA 2003a), stating that the Syngenta study cherry-picks the data by including short-term contract employees, excluding long-term employees and retirees, failing to examine actual exposure levels, and ignoring a so-called ‘healthy worker effect’ by which employed citizens should be expected to be healthier than the public as a whole.

In 2003 the EPA issued a decision allowing the continued use of atrazine in the United States with few restrictions (EPA 2003 a, b). It was then sued by the NRDC, who charged that illegal meetings between the EPA and chemical industry officials resulted in an insider-sweetheart deal on the subject. In this and a series of other lawsuits, the NRDC has charged the Bush Administration EPA with failure to act properly to protect human health and the environment from alleged deleterious effects from atrazine (NRDC v. Whitman, Northern District California 2002).

A group of documents released by the EPA on April 6, 2006 in response to these lawsuits mandates increased monitoring of watersheds, research on human health effects, and protection of endangered species. The latest chapter in this story is a letter-of-intent filed by the NRDC dated March 14 2007, charging failure by the EPA to follow through on agreements reached in the April 2006 group of documents. Of immediate concern to the NRDC are negative effects on 8 species of endangered freshwater mussels (one perhaps already extinct) in watersheds with high atrazine inputs.

The threat of atrazine in our drinking water has also gone mostly unnoticed and unstudied, but there have been residues found in public water supply in various agricultural communities in the USA (Ventura et al., 2008, Kligerman et al., 2000). People who use groundwater as their drinking water source can be exposed to atleast 0.2ppb atrazine, and atrazine may also be ingested along with corn, nuts, fruit, and wheat. Exposure safety tests conducted on mice were found to be inconclusive although DNA damage was observed (Kligerman et al., 2000).

The manager of Bourdeau Brothers Feed and Fertilizer (Sheldon, VT), Franklin county’s main supplier and custom applicator of agricultural chemicals, was interviewed about atrazine use in Vermont. Although atrazine was the most widely used herbicide used to grow corn sileage twenty years ago, its use is rare here today. According to this source it has been replaced by other herbicides that are more effective and that, unlike Atrazine, allow a field to be plowed immediately after application. One common practice is use of glyphosate in conjunction with genetically modified (“Roundup Ready”) seed, favored in part for the lack of persistant residue and low toxicity of glyphosate (D. Bourdeau, personal communication).
Conclusions/ Recommendations:

We find three areas of significant concern regarding atrazine use at current levels in the Chesapeake Bay Watershed.

First, we find reasonable evidence that current levels of atrazine found within the watershed may have significant deleterious effects on economically and culturally important fisheries and shellfish stocks. The EPA has estimated the costs of phasing out atrazine use within the United States to be about 2 billion dollars annually, mostly due to costs of alternative agricultural herbicides and non-chemical weed suppression practices (EPA, 2003a). Factoring the costs associated with eliminating or minimizing atrazine use within the Bay area should be balanced by potential economic benefits from fishery restoration.

Second, we find reasonable concern that atrazine may be contributing to the ill health of the Chesapeake Bay estuary, through direct impacts and flora and fauna production and behavior, as well as through the disruption of the normal predator prey dynamics and the trophic cascade of the ecosystem. Habitat production and water quality are also deleteriously altered by the presence of atrazine.

Third, we find reasonable concern about human health effects although we find that these potential health effects are not proven. Atrazine’s action on amphibian development involves a hormonal pathway that is well recognized in contributing to human breast and prostate cancers. In amphibians, effects are shown at environmental atrazine levels well below the EPA established maximum acceptable concentration for drinking water. High levels of human exposure to atrazine may be linked to prostate and other cancers.

In light of this evidence we find that a conservative approach to public health requires immediate action to limit agricultural use of atrazine, to reassess current standards for allowable levels in drinking water, and workplace and other exposures. An increase in monitoring the use of atrazine and its potential harm when carried by runoff within the watershed is also important in maintaining a healthy Chesapeake Bay.

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