Final_Report_Atrazine

Document Sample
Final_Report_Atrazine Powered By Docstoc
					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.
                                                                                              2


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.
                                                                                      3


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.
                                                                                            4



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).
                                                                                            5

  Figure 3: Molecular structure of
  atrazine (Stagnitti et al., 2001).          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
                                                                                            6


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.
                                                                                                7


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 (Alaska Fish Science Center, 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)
                                                                                           8


        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
                                                                                            9


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
                                                                                        10


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
                                                                                        11


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
                                                                                         12


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.
        Epifaunal communities experience most of their growth and productivity in the
summertime, when hypoxia levels are also at their highest, due to the high level of
nutrients that load the ecosystem (Sagasti et al., 2001). Eutrophia and hypoxia can have
negative impacts on these ecosystems. Crustaceans (including juvenile blue crabs) have
low tolerance to hypoxia, increased predation and stress, all of which can also reduce
benthic biomass. Juvenile blue crabs have little resistance to low oxygen levels, while
adult blue crabs can leave hypoxic regions, returning to prey on the remaining, impaired
species (Sagasti et al., 2001). Hypoxic events can be periodic or chronic, and can induce
blue crab cannibalism. Chronic hypoxia causes defined stratification between low
dissolved oxygen in the deeper waters, while the shallows remain oxygenated. Periodic
upwellings show mixing between the deep and shallow water, forming a gradient of
hypoxia increasing with depth [Figure 6]. Chronic hypoxic conditions cause adult blue
crabs populations to increase in the shallower habitats, where mortality of juveniles
increases exponentially (Eggleston et al., 2005).
                                                                                        13




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).

Endocrine Disruption in Amphibians and Concerns for Human Health:
         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
                                                                                        14


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
                                                                                         15


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
                                                                                      16


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.
                                                                                              17



Citations:

Aguilar, R., E.G. Johnson, A.H. Hines, M.A. Kramer, & M.R. Goodison. Importance of Blue
         Crab Life History for Stock Enhancement and Spatial Management of the Fishery in
         Chesapeake Bay. Reviews in Fisheries Science 16 (2008)
Alaska Fisheries Science Center. Molting: How Crabs Grow. National Marine Fisheries Service –
         NOAA Fisheries. http://www.afsc.noaa.gov/Kodiak/shellfish/cultivation/crabGrow.htm
         (accessed 2 April 2008).
Battaglin, W.A.,E.T. Furlong, M.R. Burkhardt, and C.J. Peter. Occurrence of sulfonylurea,
         sulfonamide, imidazolinone, and other herbicides in rivers, reservoirs and ground water
         in the Midwestern United States, 1998. Science Total Environment 248 (2000).
Bejarano A.C., P.L.Pennington,, M.E. DeLorenzo, and G.T.Chandler. Atrazine effects on
         meiobenthic assemblages of a modular estuarine mesocosm. Marine Pollution Bulletin 50
         (2005).
Bishop, M.J., S.L. Wear. Ecological consequences of ontogenetic shifts in predator diet:
         Seasonal constraint of a behaviorally mediated indirect interaction. Journal of
         Experimental Marine Biology and Ecology 326 (2005).
Blackmon, D.C., D.B. Eggleston. Factors influencing planktonic, post-settlement dispersal of
         early juvenile blue crabs (Callinectes sapidus Rathbun). Journal of Experimental Marine
         Biology and Ecology 257 (2001).
Bourdeau, D. Personal interview 22 April 2008.
Chapman R.N., and J.W. Stranger. Horticultural pesticide residues in water: a review of potential
         for water contamination by pesticides used in the vegetable industry in Victoria.
         Melbourne, Australia. Department of Food and Agriculture 137 (1992).
Davis, G. & B. Davis. Life History and Management of Blue Crabs. Maryland Recreational
         Fisheries. http://www.dnr.state.md.us/fisheries/recreational/articles/bluecrablhmgt.html
         (Accessed on 1 April 2008)
Delistraty, D.A., and C. Hershner. Effects of the Herbicide Atrazine on Adenine Nucleotide Levels
         in Zostera marina L. (Eelgrass). Aquatic Botany 18 (1984).
DeLorenzo, M.E., J. Lauth, P. L. Pennington, G. I. Scott and P. E. Ross. Atrazine Effects on the
         Microbial Food Web in Tidal Creek Mesocosms. Aquatic Toxicology 26 (1999).
deNoyelles F, Kettle WD, Sinn DE. The responses of phytoplankton communities in experimental
         ponds to atrazine, the most heavily used pesticides in the United States. Ecology 63
         (1982).
Dewey SL. Effects of the herbicide atrazine on aquatic insect community structure and
         emergence. Ecology 67 (1986).
Dittel, A., C. E. Epifanio, & M. Fogel. Trophic Relationships of Juvenile Blue Crabs (Callinectes
         sapidus) in estuarine habitats. Hydrobiologia 568 (2006).
Dittel, A., C.E. Epifanio, S.M. Schwalm, M.S. Fantle, and M.L. Fogel. Carbon and Nitrogen
         Sources for Juvenile Blue Crabs Callinectes sapidus in Coastal Wetlands. Marine
         Ecology Progress Series 194 (2000).
Ebersole, E.L. and V.S. Kennedy. Prey Preferences of Blue Crabs Callinectis sapidus. Feeding
         on Three Bivalves Species. Marine Ecology Progress Series 118 (1995).
Eggleston, D.B., G.W. Bell, A.D. Amavisca. Interactive effects of episodic hypoxia and
         cannibalism on juvenile blue crab mortality. Journal of Experimental Marine Biology
         and Ecolgy 325 (2005).
Environmental Protection Agency. (2003a). Atrazine Interim Reregistration Decision (IRED).
         January 2003.
                                                                                               18

Environmental Protection Agency. (2003b). Revised Atrazine Interim Reregistration Decision
        (IRED). October 31, 2003
Environmental Protection Agency. (2006). Finalization of Atrazine IRED, and Completion of
        Tolerance Reassessment and Reregistration Eligiblity Process. April 6, 2006.
Epifanio, C.E., A.K. Masse and R.W. Garvine. Transport of blue crab larvae by surface currents
        off Delaware Bay, USA. Marine Ecology Progress Series 54 (1989).
Flinders, C.A., R.J. Horwitz and T. Belton. Relationship of fish and macroinvertebrate
        communities in the mid-Atlantic uplands: Implications for integrated assessments.
        Ecological Indicators 8-5 (2008).
Fortin, M.G., C.M. Couillard, J. Pellerin, M. Lebeuf. Effects of salinity on sublethal toxicity of
        atrazine to mummichog (Fundulus heteroclitus) larvae. Marine Environmental Research
        65 (2008).
Gonzales-Muria C., A. Munoz-Rueda, F. Hernando, and M. Sanchez-Diaz. Effect of Atrazine and
        Methabenzthiazuron on Oxygen Evolution and Cell Growth of Chlorella pyrenoidosa.
        Weed Research 25 (1985).
G. S. Ward, Ballantine, L., Acute and chronic toxicity of atrazine to estuarine fauna. Estuaries
        Vol. 8, No. 1, p. 22-27 March 1985.
Jongen, V.H.W.M, Jhhh Thijssen, H. Hollema, G.H. Donker, J.G. Santema, A.J.G. Heineman.
        International Journal of Gynecological Cancer (2005).
Hall, L.W., M. C. Ziegenfuss, R. D. Anderson, T. D. Spittler and H. C. Leichtweis. Influence of
        salinity on atrazine toxicity to a Chesapeake Bay copepod (Eurytemora affinis) and fish
        (Cyprinodon variegates). Estuaries 17 (1994).
Hamilton PB, Jackson G, Kaushik NK, Solomon KR. The impact of atrazine on lake periphyton
        communities including carbon uptake dynamics using track autoradiography.
        Environmental Pollution 46 (1987).
Hayes TB, Collins A, Mendoza M, Noriega N, Stuart AA, Vonk A. Hermaphroditic,
        demasculinized frogs exposed to the herbicide atrazine at low ecologically relevant
        doses. Proceedings of the National Acadamy of Science 99,(2002).
Hayes, T.B., Haston K, Tsui M, Hoang A, Haeffele C, and Vonk A. Atrazine-induced
        Hermaphroditism at 0.1 ppb in American Leopard Frofs (Rana pipiens): Laboratory and
        Field Evidence. Environmental Health Perspectives 10 (2002).
Hayes T.B., Haston K, Tsui M, Hoang A, Haeffele C, and Vonk A. Atrazine-Induced
        Hermaphroditism at 0.1 ppb in American Leopard Frogs. Environmental Health
        Perspectives 111 (2003).
AddedHines, A. H, E.G. Johnson, A.C. Young, R. Aguilar, M.A. Kramer, M. Goodison, O.
        Zmora, &Y. Zohar. The Chesapeake Bay Blue Crab (Callinectes sapidus): A
        Multidisciplinary Approach to Responsible Stock Replenishment. Reviews in Fisheries
        Science. 16 (2008).
Hines, A. Fish and Invertebrate Ecology. Smithsonian Environmental Research Center.
        http://serc.si.edu/labs/fish_invert_ecology/index.jsp (accessed 4 April 2008)
Hoeinghaus, D.J. & S. E. Davis III. Size-based Trophic Shifts of Saltmarsh Dwelling Blue Crabs
        Elucidated by Dual Stable C and N Isotope Analyses. Marine Ecology Progress Series.
        334 (2007).
Hovel, K.A., N.L. Romuald. Effects of seagrass habitat fragmentation on juvenile blue crab
        survival and abundance. Journal of Experimental Marine Biology and Ecology 271
        (2002).
Hovel, K.A. and M.S. Fonseca. Influence of Seagrass Landscape Structure on the Juvenile Blue
        Crab Habitat Survival Function. Marine Ecology Progress Series. 300 (2005).
Johnson, J.R., and K.T. Bird. The Effects of the Herbicide Atrazine on Ruppia maritima L.
        Growing in Autotrophic Versus Heterotrophic Cultures. Botanica Marina 38 (1995).
                                                                                             19

Jones, M.B. and C.E. Epifanio. Settlement of brachyuran megalopae in Delaware Bay: an
        analysis of time series data. Marine Ecology Progress Series. 125 (1995).
Jones, T.W. and L. Winchell. Uptake and photosynthetic inhibition by atrazine and its
        degradation products on four species of submerged vascular plants. Journal of
        Environmental Quality 13 (1984).
King, R.S., A.H. Hines, F.D. Craige, S. Grap. Regional, watershed and local correlates of blue
        crab and bivalve abundances in subestuaries of Chesapeake Bay, USA. Journal of
        Experimental Marine Biology and Ecology 319 (2005).
Kuhlman, M.L. and A.H. Hines. Density-dependent predation by Blue Crabs Callinectis sapidus
        on Natural Prey Populations of Infaunal Bivalves. Marine Ecology Progress Series. 295
        (2005).
Marshall, H.G. and R.W. Alden. Comparison of Phytoplankton Assemblages in the Chesapeake
        and Delaware Estuaries (USA), with Emphasis on Diatoms. Hydrobiologia 269/270
        (1993).
Marshall, H.G. and K. K. Nesius. Phytoplankton Composition in Relation to Primary Production
        in Chesapeake Bay. Marine Biology. 125 (1996).
McEnerney, J.T., and D.E. Davis. Metabolic Fate of Atrazine in the Spartina alterniflora-
        Detritus-Uca pugnax Food Chain. Journal of Environmental Quality. 8 (1979).
Moerke, A.H., Lamberti, G.A. Scale-dependent influences on water quality, habitat, and fish
        communities in the streams of the Kalamazoo River Basin, Michigan (USA). Aquatic
        Science 68 (2006).
Mokbel K. Evolving role of aromatase inhibitors in breast cancer. International Journal of
        Clinical Ontology 7 (2002).
Moksnes, P.O. & K. L. Heck. Relative Importance of Habitat Selection and Predation for the
        Distribution of Blue Crab Megalopae and Young Juveniles. Marine Ecology Progress
        Series. 308 (2006).
Orth, R.J. and J. van Montfrans. Habitat Quality and Prey Size as Determinants of Survival in
        Post-Larval and Early Juvenile Instars of the Blue Crab Callinectes sapidus. Marine
        Ecology Progress Series. 260 (2002).
Pardieck, R.A., R.J. Orth, R.J. Diaz, R.N. Lipcius. Ontogenetic Changes in Habitat Use by Post-
        larvae and Young Juveniles of the Blue Crab. Marine Ecology Progress Series. 186
        (1999).
Pollehne, F., G. Jost, E. Kerstan, B. Meyer-Harms, M. Reckermann, M. Nausch and D. Wodarg.
        Triazine Herbicides and Primary Pelagic Interactions in an Estuarine Summer Situation.
        Journal of Experimental Marine Biology and Ecology. 238 (1999).
Posey, M.H., T.D. Alphin, H. Harwell, B. Allen. Importance of low salinity areas for juvenile
        blue crabs, Callinectes sapidus Rathbun, in river-dominated estuaries of southeastern
        United States. Journal of Experimental Marine Biology and Ecology 319 (2005).
Rugolo, L.J., K.S. Knotts, , A.M. Lange, and V.A. Crecco. Stock assessment of Chesapeake Bay
        blue crab (Callinectes sapidus Rathbun). Journal of Shellfish Restoration 17 (1998).
Ryer, C.H., J. Van Montfrans, and K.E. Moody. Cannibalism, Refugia, and the Molting Blue
        Crab. Marine Ecology Progress Series. 147 (1997).
Sagasti, A., L.C. Schaffner, J.E. Duffy. Effects of periodic hypoxia on mortality, feeding and
        predation in an estuarine epifaunal community. Journal of Experimental Marine Biology
        and Ecology 258 (2001).
Seitz, R.D., R.N. Lipcius, M.S. Seebo. Food availability and growth of the blue crab in seagrass
        and unvegetated nurseries of Chesapeake Bay. Journal of Experimental Marine Biology
        and Ecology 319 (2005).
Solomon, K.R., D.B. Naker, R.P. Richards, K.R. Dixon, S.J., Klaine, T.W. LaPoint, R.J. Kendall,
        C.P. Weisskopf, J.M. Giddings, J.P. Giesy, L.W. Hall Jr., and W.M. Williams. Ecological
                                                                                              20

         Risk Assessment of Atrazine in North American Surface Waters. Environmental
         Toxicology and Chemistry. 15 (1996).
Stagnitti, F., M. Graymore, G. Allinson. Impacts of atrazine in aquatic ecosystems.
         Environmental International 26 (2001).
Steinberg CEW, Lorenz R, Spieser O.H. Effects of atrazine on swimming behavior of zebrafish,
         Brachydanio rerio. Water Restoration 29 (1995).
Stevenson, J.C., T.W. Jones, W.M. Kemp, W.R. Boynton and J. C. Means. An Overview of
         Atrazine Dynamics in Estuarine Ecosystems. In: Proceedings of the Workshop on
         Agrochemicals and Estuarine Productivity, Beaufort, North Carolina, September 18-19,
         1980 (1982).
Tang, J.X., K.D. Hoagland, and B.D. Siegfried. Differential Toxicity of Atrazine to Selected
         Freshwater Algae. Bulletin of Environmental Contamination and Toxicology. 59 (1997).
Tilburg, C.E., A.I. Dittel, and C.E. Epifanio. Retention of crab larvae in a coastal null zone.
         Estuarine Coastal & Shelf Science. 72 (2007).
United States Geological Survey. Chesapeake Bay Monitoring Program.
         http://va.water.usgs.gov/chesbay/RIMP/generalinfo.html. (accessed on 3 April 2008)
Van Dijk, H.F.G., and Guichert, R. Water Soil Air Pollution 115 (1995).
Van Montfrans J., C.H. Ryer, and R. J. Orth. Substrate Selection by Blue Crab Callinectes
         sapidus megalopae and First Juvenile Instars. Marine Ecology Progress Series. 260
         (2003).
Ventura, B. de Campos, D. de Fransceschi de Angelis, M.A. Marin-Morales. Mutagenic and
         genotoxic effects of the Atrazine herbicide in Oreochromis niloticus (Perciformes,
         Cichlidae) detected by the micronuclei test and the comet assay. Pesticide Biochemistry
         and Physiology 90 (2008).
Vervliet-Scheebaum, M., R. Ritzenthaler, J. Normann, E. Wagner. Short-term effects of
         benzalkonium chloride and atrazine on Elodea canadensis using a miniaturised
         microbioreactor system for an online monitoring of physiologic parameters.
         Ecotoxicology and Environmental Safety 69 (2008).
Yeh, H.J., C.Y. Chen. Toxicity assessment of pesticides to Pseudokirchneriella subcapitata
         under air-tight test environment. Journal of Hazardous Materials A131 (2006).
Zinski, S. Blue Crab Growth and Molting. The Blue Crab Archives.
         http://www.bluecrab.info/molting.html (accessed on 1 April 2008)
Vervliet-Scheebaum, M., R. Ritzenthaler, J. Normann, E. Wagner. Short-term effects of
         benzalkonium chloride and atrazine on Elodea canadensis using a miniaturised
         microbioreactor system for an online monitoring of physiologic parameters.
         Ecotoxicology and Environmental Safety 69 (2008).
Yeh, H.J., C.Y. Chen. Toxicity assessment of pesticides to Pseudokirchneriella subcapitata
         under air-tight test environment. Journal of Hazardous Materials A131 (2006).
Zinski, S. Blue Crab Growth and Molting. The Blue Crab Archives.
         http://www.bluecrab.info/molting.html (accessed on 1 April 2008)

				
DOCUMENT INFO