CARBARYL EFFECTS ON AMPHIBIAN SPECIES WITH IMPLICATIONS FOR FURTHER RESEARCH

A Literature Review

By Laura Mills

Southeast Missouri State University Spring 2008

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Abstract. Declines in amphibian populations have been reported worldwide. Increased pesticide use is hypothesized to contribute to the loss of amphibian species. A comprehensive review of literature discusses water contamination by the insecticide, carbaryl, to illustrate the challenges of ecotoxicology and the study of amphibian declines. Because amphibian survival is determined by complex interactions of multiple abiotic and biotic factors, it is logical to conclude that better understanding of the declines will be found through the examination of the interactive effects of multiple stressors. In addition to long-term research and monitoring, further study of multiple stressors could provide data necessary to slow the rate of amphibian declines and may serve as a model for addressing the global issue of decreasing biodiversity. Introduction

Brief History of Amphibian Declines

Amphibian populations have been declining worldwide since the 1970s in a trend

believed to be too dramatic and extensive to be attributed to natural demographic

variations (Stuart et al. 2004). The estimated rate of extinction currently surpasses

fluctuations recognized over the last 100,000 years (Eldridge 1998). Thirty-four species

of amphibians are reported to have vanished between the years 1500 and 2004. Nine of

those extinctions occurred between 1980 and 2004, and an additional 113 species have

apparently disappeared since 1980, but have not been formally classified as “extinct”

(Stuart et al. 2004).

The loss of amphibian populations became a global concern in 1989, at the First

World Congress of Herpetology (Collins and Storfer 2003). In 1990, a U.S. National

Research Council Workshop began to investigate the phenomenon (Wake 1991). By

1993 over 500 different populations of frogs and salamanders were recognized as

declining or listed as species of special conservation concern (Alford and Richards 1999).

At the Third World Congress of Herpetology in 1997, the critical need for further

research to address the increasing threat of potential amphibian extinction was identified

(Collins and Storfer, 2003).

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In 2004, the Global Amphibian Assessment (GAA) of the World Conservation

Union (IUCN), reported that approximately 32.5% of amphibian species were globally

threatened, 7.4% were critically endangered, and 22.5% were not assessed due to lack of

data. At least 43.2% of amphibian species were experiencing population decrease, while

0.5% was increasing, 27.2% were stable, and 29.1% have an unknown trend (Stuart et al.

2004).

Significance of Amphibian Declines

Loss of biodiversity is a global environmental concern (Wake 1991, Blaustein et

al. 1994a). It is important to understand amphibian declines because it could serve as a

model for addressing the general issue of decreasing global biodiversity (Houlahan et al.

2000, Alford et al. 2001). Amphibians are considered biomarkers of environmental stress

(Blaustein et al. 1994, Blaustein and Wake 1995), and the severe decline in amphibian

biodiversity could be an indication of serious environmental problems often associated

with human activities (Wake 1991). Amphibian losses are occurring simultaneously,

expansively, and rapidly. Most notably, population declines are occurring in protected

natural areas, so efforts to protect species have not been effective (Collins and Storfer

2003).

Amphibians provide ample biomass and biodiversity to their environment

(Blaustein et al. 1994a, Boyer and Grue 1995). The species have important roles in the

food web. Amphibians are predators, as well as, prey for other trophic levels (Blaustein

et al. 1994a, Wake 1991) and they may be herbivorous, carnivorous, or omnivorous,

depending on species and life stage (Zug 1993). The ecological impact of loss of

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amphibian biodiversity is difficult to forecast; but, because of their role in trophic

dynamics, the worldwide reduction of amphibian species and subsequent alteration of the

biological community could have significant consequences for other organisms

(Blaustein et al. 1994a). The importance of amphibian populations may not be fully

recognized until they have vanished, and the consequences could be immense and

irreversible.

Major Hypotheses

Six major hypotheses to explain the cause of the amphibian population declines

are introduced species, over-exploitation, land alteration, global change, disease, and the

increased use of pesticides and other toxic chemicals. Studies investigating amphibian

population declines demonstrate that the underlying mechanisms are complex and

involve interactions among abiotic and biotic components (Blaustein and Kiesecker

2002). Many researchers agree that, generally, amphibian declines are not likely to be

the result of one single factor, but rather, the product of synergistic interactions of

multiple stressors (Alford and Richards 1999, Blaustein and Kiesecker 2002).

Amphibian reaction to one stressor can be different when affected by the presence of

another stressor (Kiesecker et al. 2001a). Therefore, studying how these factors interact

should provide better understanding of the reasons for increasing rates of amphibian loss

(Blaustein and Kiesecker 2002). It is especially important to examine human impacts on

amphibian declines so that measures can be taken to slow the rate of extinctions (Collins

and Storfer 2003).

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Introduced Species

The introduction of non-native species can have negative consequences for

amphibian populations. For example, mass introductions of predatory fish have caused

an increase in amphibian exposure to predators which they have not had previous

interactions with and therefore have not evolved anti-predator mechanisms against

(Alford and Richards 1999). Amphibian response mechanisms that would protect them

against natural predators could thereby be ineffective against the alien species (Gamradt

and Kats 1996). When predatory fish are introduced to normally fish-free waters, rapid

extinction of amphibian assemblages can occur (Fisher and Shaffer 1996). Predation, a

direct effect, as well as indirect effects including competition, hybridization, and the

introduction of pathogens, cause devastation to native amphibian populations as a result

of the introduction of alien species to a habitat (Collins and Storfer 2003).

Over-exploitation

Ranid species are at the highest risk of over-exploitation due to extensive

harvesting for frog legs for human consumption (Stuart et al. 2004). Over-exploitation

has caused rapid decline in at least 50 amphibian species and is considered a major

source of amphibian declines in East and Southeast Asia (Stuart et al. 2004). In the past,

the frog leg trade has accounted for annual exports of approximately two hundred million

frogs in Asia (until 1995) and seventy million frogs in India (by 1990) (Oza 1990).

Lannoo et al. (1994) reported that harvesting for frog legs accounted for at least one third

of the decline in amphibian species in one Iowa County where estimates of frog

abundance decreased from at least 20 million frogs in 1920 to about 50,000 in 1992.

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Land-use/land-cover change

Alford and Richards (1999) asserted that “habitat modification is the best

documented cause of amphibian population declines.” Meyer and Turner (1992) divided

this hypothesis into two sub-categories: land-use change and land-cover change. A

change in the way people use the land is termed land-use change, while land-cover

change is the alteration of the “physical or biotic character of the land surface,” (Meyer

and Turner 1992). Land-use change is basically disturbance of the environment; for

example, the introduction of non-native species or pollution of the water, which has

various adverse effects on amphibian species (see Increased Use of Pesticides and Other

Chemicals below). Land-cover change typically involves habitat loss or food scarcity;

for example, wetland drainage, deforestation, and agricultural intensification. Almost

120,000 ha of wetlands are lost each year (Boyer and Grue 1995). Petranka (1998)

declared that the biggest threat facing the smallmouth salamander, Ambystoma texanum,

is the conversion of such bottomland habitats to agricultural areas. High-traffic

roadways, which may have considerable impacts on anuran populations, are another

example of land-cover change (Lamboureux and Madison 1999). Land-use change and

land-cover change are noted to cause of global loss of biodiversity, including the loss of

amphibian species (Collins and Storfer 2003).

Global Change

The global change hypothesis encompasses various changes in the planet. Global

climate change as a result of depletion of stratospheric ozone is one concern. For

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example, certain species of amphibians require fishless ponds for breeding and thus

depend on ephemeral pools that cannot sustain populations of fish. Altered precipitation

patterns, as observed in the neotropics, can cause ponds to fill later and dry earlier,

resulting in increased density in the limited breeding sites (Donnelly and Crump 1998).

Density mediated effects of increased competition and predation may then induce shifts

in amphibian distributions (Donnelly and Crump 1998).

Stratospheric ozone depletion is also a source of increased ultraviolet-B (UV-B)

radiation (290 – 320 nm) at the terrestrial surface (McKenzie et al. 2007), which could

potentially affect amphibian populations in certain locations. Blaustein et al. (1994b)

measured levels of photolyase, a photoreactivating DNA repair enzyme that mends UV-B

damage, in eggs and oocytes of ten amphibian species (Xenopus laevis, Rana cascadae,

Hyla regilla, Bufo boreas, Taricha granulosa, Ambystoma gracile, Ambystoma

macrodactylum, Plethodon dunni, Plethodon vehiculum, and Rhyacotriton variegatus),

and performed field experiments to measure the effects of UV-B on hatching success of

three anuran species (Rana cascadae, Bufo boreas, and Hyla regilla). The study found

the variation in levels of photolyase among species to be correlated with exposure to

sunlight of natural egg laying locations. Furthermore, Blaustein et al. (1994b) found a

remarkable correlation between the population status of the species tested for hatching

success, and their ability to repair UV-B damage: the species experiencing drastic

population declines (i.e. Rana cascadae and Bufo boreas) were those with the lowest

photolyase activity.

Note that, with the exception of laboratory-reared Xenopus laevis, all amphibian

subjects were collected from mountainous populations, which may be more at risk for

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exposure to high levels of UV-B irradiance than populations at lower elevations. Kerr

and McElroy (1993) reported increases in UV-B radiation (35% per year in winter, and

7% per year, in summer for 4 years) associated with detections of decreasing ozone at the

same latitudes (44ºN) in Canada as the field sites used by Blaustein et al. (1994b). Most

importantly, Blaustein et al. (1994b) related adverse effects of UV-B in spring field trials

to the observation by Kerr and McElroy (1993) that rising levels of UV-B in late spring

may have a much greater impact on some species if they occur at critical stages of

development.

Embryos may be at risk for lethal and sublethal effects resulting from exposure to

UV-B radiation. Amphibian larvae and adults are generally able to seek refuge from

ambient UV-B, while embryos may be exposed to continuous levels of sunlight because

they are stationary and must endure the environmental conditions in the location in which

they are deposited (Zaga et al. 1998). Anzalone et al. (1998) observed high rates of

embryonic mortality in Hyla cadaverina and Taricha torosa from exposure to solar UV-

B radiation at natural breeding sites in California (34ºN latitude, 290 m altitude), while

protection from solar UV-B resulted in significantly greater hatching success.

Amphibian eggs that are exposed to increased ambient levels of UV-B radiation have

decreased hatching success and suffer sublethal effects that alter growth and

development, behavior, anatomy, and physiology (Blaustein et al. 2001b). Constant

mortality in early amphibian life stages may eventually lead to declining population

(Blaustein and Kiesecker 2002).

Disease

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Emerging Infectious Diseases (EIDs) are diseases that have recently increased in

incidence or geographical range, have impacted new host populations or increased in

pathogenicity, and developed from newly discovered or newly evolved pathogens

(Daszak et al. 2003). Three pathogens inducing EIDs that are believed to contribute to

amphibian declines are Batrachochytrium dendrobatidis, Saprolegnia ferax, and

Ambystoma tigrinum virus (ATV) (Blaustein and Kiesecker 2002).

Batrachochytrium dendrobatidis is a fungus responsible for the disease,

Chytridiomycosis, which has been linked to mass die-offs in numerous amphibian species

around the world, especially in Latin America (Lips et al. 2006). Increasing evidence

suggests that Batrachochtrium dendrobatidis is a recently evolved pathogen contributing

to amphibian declines (Lips et al. 2006, Daszak et al. 2003, Morehouse et al. 2003,

Weldon et al. 2004). Furthermore, studies suggest that amphibian susceptibility to the

disease may be influenced by global climate change (Berger et al. 2004, Pounds et al.

2006, Bosch et al. 2007).

Saprolenia ferax, a pathogenic oomycete, has contributed to significant

embryonic mortality and has been proposed to be a contributing factor in the declines of

the boreal toad (Bufo boreas) and the Cascades frog (Rana cascadae) in the Pacific

Northwest United States (Daszak et al. 2003). Kiesecker et al. (2001a) showed that UV-

B radiation has a synergistic effect on boreal toads in the presence of the pathogen.

Increased mortality was observed in Saprolegnia-infected toads exposed to increased

levels of UV-B radiation (Kiesecker et al. 2001a). Other factors may influence

amphibian susceptibility to the pathogen as well. Kiesecker et al. (2001b), for instance,

found that introduced fishes are a primary source of Saprolegnia infection. The disease

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may be directly transmitted to amphibian embryos exposed to infected fish or indirectly

transmitted by soil substrate carrying the pathogen (Kiesecker et al. 2001b).

The iridovirus, Ambystoma tigrinum virus (ATV) can also be transmitted directly

or indirectly by salamanders exposed to ATV (Brunner et al. 2007). Cofactors may also

play a role in the spread of the ATV (Blaustein and Kiesecker, 2002). For example,

Rojas et al. (2005) observed that temperature affected percent mortality and time to death

of salamanders exposed to ATV. When exposed at 26º C, most of the salamanders

survived, but at 18º C all of the salamanders died, and at 10º C almost all of the

salamanders died as well (Rojas et al. 2005). The effects observed in this study could

provide reason for cyclic viral epizootics of A. tigrinum in nature which could be

influenced by seasonal variations in water temperatures (Rojas et al. 2005). Exposure to

the widely used herbicide, atrazine may also influence salamander susceptibility to ATV

and mortality and contribute to ATV epizootics (Forson and Storfer 2006a, b)

While pathogens may be present in healthy animals, a weakened immune system

allows the disease to take effect (Alford and Richards 1999). Therefore, the sublethal

effects of other environmental stressors could make amphibians more susceptible to

disease (Carey 1993, Kiesecker 2002). Of particular interest to researchers are the

impacts of global change on amphibian susceptibility to and mortality by disease

(Blaustein and Kiesecker 2002).

Increased Use of Pesticides and Other Chemicals

Pollutants are able to stress amphibians in a variety of ways (Blaustein and

Kiesecker 2002). They can cause mortality, change behavior, decrease growth rates,

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disrupt endocrine functions, and suppress the immune system (Alford and Richards

1999). Studies have revealed that amphibian species are as sensitive, if not more

sensitive, than other aquatic organisms when exposed to certain contaminants (Boyer and

Grue 1995, Blaustein and Wake 1995). This is likely because they are susceptible to

dermal absorption of toxicants in the water at all life stages (Boyer and Grue 1995).

Amphibians respire through a moist, permeable skin and lay unshelled eggs that are

directly exposed to soil, water, and sunlight, and easily absorb environmental

contaminants (Blaustein and Kiesecker 2002, Blaustein et al. 2003). Considerable

disturbance of the aquatic ecosystem can result from chemical contamination (Relyea and

Hoverman 2006). Owing to the physiological traits and central position in the food

chain, the effects of contaminants on amphibians are expected to be severe (Broomhall

2005).

According to Ongley (1996), agriculture is the number one supplier of water

pollution worldwide. Chemical fertilizers and pesticides are the leading source of water

contamination by agriculture (Horne 2001). The use of chemicals in agriculture has

increased drastically since the late 1940s (Boyer and Grue 1995), and with the

biotechnological development of crops resistant to pesticide applications, the use of

agrichemicals is growing rapidly. In the first eight years that genetically modified crops

were available to farmers, the use of pesticide resistant corn, soybean and cotton crops

led to a 122 million pound increase in pesticide use (Benbrook 2004).

Pesticides enter aquatic systems through direct application, terrestrial runoff,

erosion, and aerial drift (LeNoir et al. 1999, Relyea and Hoverman 2006). Pesticides may

even affect amphibians in remote and considerably undisturbed habitats through

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atmospheric transport (Blaustein et al. 2003). Many amphibian species inhabit very small

ephemeral pools where even minute amounts of pesticide can result in significant

chemical concentrations (Sanders 1970, Blaustein et al. 2003). In addition, breeding and

larval development are periods when amphibian species are generally restricted to their

aquatic environment. These critical periods in amphibian life histories occur during the

warmer seasons of the year, typically during growing seasons when agrichemicals are

applied to the land. Because species remain in their aquatic habitat for months or even

years, they may be exposed to contaminants numerous times (Boone et al. 2001) due to

repeated applications of agrichemicals. For instance, carbaryl is recommended by one

manufacturer (Ortho) to be reapplied every 7 to 10 days, or as needed (Boone et al.

2001). Thus, amphibian species may be quite vulnerable to agricultural contaminants

(Blaustein and Kiesecker 2002).

Lack of Protection against Water Contamination

The need for increased protection of amphibians and other aquatic species

through the development of water quality criteria has been recognized by researchers

(Boyer and Grue 1995). Though the number of ecotoxicology studies published annually

has increased tremendously since 1992 (Relyea and Hovermann 2006), there is little data

to establish how contaminants affect amphibian populations in the environment

(Blaustein and Kiesecker 2003).

Limitations in Testing

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There are thousands of different pesticides used around the world (Relyea and

Hovermann 2006), not to mention countless other contaminants that enter aquatic

systems (Koplin et al. 2002). As a requirement under the Toxic Substances Control Act,

toxicity tests must be conducted to determine the hazardous effects of various chemicals

(Relyea and Mills 2001). Due to financial and time limitations, however, acute toxicity

analyses employing only a few model organisms are often used to determine acceptable

levels of a chemical concentration that is safe for humans and non-target organisms

(Cooney, 1995).

LC50 tests are the most commonly utilized of these assessments (Relyea and

Mills 2000). An LC50 value is an estimate of the lethal concentration of a chemical

necessary to kill 50% of a test population. The estimate is used to determine the

maximum amount of a toxicant that can be tolerated by non-target species and to set

environmental safety standards; however, because of short exposure periods (and

generally high chemical concentrations) used these values may be overestimated (Relyea

and Mills 2000), (see LC50 analyses below).

Furthermore, acute toxicity tests may detect the toxicity that directly affects a

species, but the results of these experiments do not accurately reflect the complex effects

of chemical contamination in an actual aquatic community (e.g., sublethal effects,

indirect effects, additive effects) (Blaustein and Kiesecker 2002). Various biotic (e.g.,

predators, competitors, food resources) and abiotic (e.g., temperature, pH, UV-radiation)

factors that are present in the environment can interact with the amphibian, with the

chemical, or with other factors to produce effects that cannot be predicted in single-factor

studies (Saura-Mas et al. 2002).

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Moreover, pesticide registration does not require amphibian tests, thus

experiments with fish and aquatic invertebrates have generally determined the tolerance

level for aquatic chemical exposure of amphibian species (Bridges et al. 2002). Because

fish and other aquatic invertebrates have different sensitivities to toxicants than

amphibians, the evaluation of aquatic chemical exposure to amphibians is of extreme

importance in regulating agrichemicals (Bridges et al. 2002). Amphibian species are

frequently, but not always, more sensitive to water contamination than other vertebrate

species, and sensitivities can differ depending on the contaminant being tested (Bridges et

al. 2002). Also, sensitivity to toxicants can vary among species (Bridges and Semlitsch

2000), among populations of a single species (Bridges and Semlitsch 2000) and among

life stages (Widder and Bidwell 2006). In addition, the sensitivities of various species

must not be determined solely by the results of acute toxicity tests, but with additional

studies including chronic exposure and sublethal concentrations to gain more accurate

results on species’ sensitivities (Bridges et al. 2002).

Limitations in Interpretations

Pesticide concentrations detected in aquatic systems are frequently lower than

those used in toxicity experiments. However, environmental concentrations are generally

considered “snapshots in time” rather than detections of maximum concentrations (i.e. at

the time of application) in an aquatic system (Relyea and Hovermann 2006). The

accumulation and the rate of breakdown of a chemical within a body of water can be

affected by various factors including the size of the water body, timing of application,

pH, temperature, sunlight exposure and other factors. Koplin et al. (2002) states that

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astonishingly little is certain about the degree of environmental occurrence, transport, and

ultimate fate of many synthetic organic chemicals after their intended use. Lack of

extensive data on pesticide concentrations in the environment thus restrict the

conclusions that can be drawn from toxicity studies (i.e. the relevance of the

concentrations used may not be clear) (Relyea and Hovermann 2006).

Agrichemicals (and other contaminants) that are not intended for or legally

registered for application to aquatic systems still appear in these environments (e.g.

Thompson et al. 2004, Koplin et al. 2002) as a consequence of runoff, areal drift, erosion,

direct overspray, etc. (Peterson et al. 1994). Until we know how these agrichemicals are

affecting amphibian species, we cannot confirm that they are not having serious adverse

effects on amphibian species and the ecosystem. Hence, further research is needed to

determine chemical concentrations of pesticides in aquatic systems and the effects of

these chemicals on non-target organisms to better understand the threat the contaminants

pose to amphibian species (Relyea and Hovermann 2006).

Amphibian Declines near Areas of Intense Cultivation

A relationship exists between amphibian declines and proximity to agricultural

lands. Pesticides are carried downwind and deposited in amphibian habitats. For

example, vast quantities of agrichemicals are applied to the San Joaquin Valley of

California each year (Sparling et al. 2001). The valley is one of the most highly

cultivated and productive regions in the world, and relies on the use of millions of

kilograms of pesticides, particularly organophosphate (OP) pesticides (Aston and Seiber

1997). Through volatization and airborne transport by dominant wind patterns, the

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pollution is carried from the San Joaquin Valley and eventually deposited in the nearby

Sierra Nevada mountains (Zabik and Seiber 1993, Aston and Seiber 1997, Le Noir et al.

1999). Zabik and Seiber (1993) showed that considerable amounts of OP insecticides (as

high as 100 ng/L for diazinon, 10 ng/L for chlorpyrifos, and 10 ng/L for paration) used in

the San Joaquin Valley are transported to the Sierra (e.g., Sequoia National Park).

Between 50% and 100% of the precipitation and water samples taken from different sites

(i.e. at 533 m elevation and at 1,290 m elevation in Sequoia National Park and at 2,200 m

elevation in the Lake Tahoe Basin) within the Sierra Nevada mountains, contained OPs

and other currently used pesticides (as high as 85 ng/L chlorothalonil, 13 ng/L

chlopryrifos, 24 ng/L malathion, 19 ng/L diazinon, and 2 ng/L trifluralin,

hexachlorocyclohexane, and endosulfan) (McConnell et al. 1998). An estimated annual

loading of chlorpyrifos, an organophosphate, was determined to be 24 to 31 kg/year for

the Sequoia National Park area (McConnel et al. 1998). Concentrations in air and water

correlated roughly with peak pesticide application periods in the Valley (McConnel et al.

1998). The detected concentrations are less than acute toxicity values of amphibian

species, though additive, synergistic, and indirect effects of the pesticides must be

considered (LeNoir et al. 1999).

The most severe declines in amphibian species in California have been found in

the Sierra Nevada Mountains (Drost and Fellers 1996, Fellers and Drost 1993). Air

masses laden with ozone, aerosols, and agrichemicals are moved into the Sierra causing

deteriorated air quality in the foothills and the mountains (Aston and Seiber 1997).

Widespread ozone damage has occurred throughout the Sierra (i.e., possible increased

UV radiation effects) and there is concern that agrichemical effects may be in part

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responsible for declining number of native amphibians (Boyer and Grue 1995). In some

areas there have been reports that no frogs can be found, though only 10 yr earlier there

were well documented thriving populations (Aston and Seiber 1997).

Sparling et al. (2001) measured cholinesterase activity and residues in a

California anuran species, Hyla regilla, and found that the frequency of detections of

pesticide residues, as well as concentrations of pesticides in amphibian tissues, correlated

with patterns of consistent agrichemical drift and areas of amphibian declines.

Cholinesterase-inhibiting pesticide exposure could therefore be responsible for sharp

population declines in frog species in Sierra Nevada range in California (Sparling 2001,

Davidson 2004).

Carbaryl

Background

Subsequent to the prohibition of organochlorine (OC) pesticides in the 1970s, the

use of organophosphate (OP) and carbamate pesticides increased dramatically (Sparling

et al. 2000). The increase in OP and carbamate pesticide use immediately preceded or

coincided with amphibian declines, which are believed to have begun in the early to mid-

1970s in the Western United States (Jennings 1996). Carbaryl (1-naphthyl-N-

methylcarbamate) is a broad-spectrum insecticide, better known by the trade name

SEVIN, that is common in agriculture, forestry, lawn and garden use throughout the

United States and Canada (Rohr et al. 2003). It is a neurotoxin that kills amphibian

species by inhibiting acetylcholinesterase (Relyea 2005) which results in the

accumulation of acetlycholine in the synapse, causing the nerve to repeatedly fire (Zaga

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et al. 1998). Examining the effects of carbaryl on amphibian species is important because

cholinesterase-inhibiting pesticides are most strongly associated with amphibian declines

and carbaryl may serve as a model chemical for understanding the effects of carbamate

and organophosphates (i.e., cholineseterase inhibitors) (Cox 1993, Rand 1995).

Concentration and Breakdown in the Environment

Carbaryl is one of the most frequently detected pesticides in urban streams

(Phillips et al. 2007, Schreder and Dickey 2005). Environment Canada calculated the

Expected Environmental Concentration (EEC), the expected concentration resulting from

maximum proposed application rate to a 15 cm deep water body, to be 3.667 mg/L

(Peterson et al. 1994). Past studies have found the insecticide at concentrations as high as

4.8 mg/L (Norris et al. 1983), although detections of concentrations greater than 0.01

mg/L have not been reported recently (Brena et al. 2005, Koplin et al. 2002, Phillips et al.

2007).

It is possible that greater concentrations are present in the environment at levels

that may have adverse effects on amphibian species (see Limitations in Interpretations

above). For instance, levels of the insecticide have been determined in larger water

bodies where the significant levels of carbaryl are less likely to exist when compared to

shallow waters where smaller amounts of the toxicant result in higher concentrations.

Also, carbaryl decomposes rather quickly under most environmental conditions (Sparling

et al. 2001); therefore peak concentrations of the contaminant have not necessarily been

detected (Relyea and Hovermann 2006). Studies on the concentration of carbaryl in

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small ephemeral pools receiving direct overspray could provide a “worst-case scenario”

for the insecticide concentrations in the environment.

Although recent detections of carbaryl in the environment are at low

concentrations, frequent detections of high levels of 1-naphthol, the major breakdown

product of carbaryl, in groundwater and surface water have been observed (Brena et al.

2005). A study conducted by Brena et al. (2005), for example, did not detect significant

levels of carbaryl, but considerable amounts of 1-naphthol were detected more often than

any other contaminant. Brena et al. (2005) suggests that the breakdown of carbaryl

occurred prior to sampling, probably catalyzed by enzymes from soil microorganisms;

though the detection of 1-naphthol may serve as an indicator that short-lived

agrochemicals also reach the groundwater. 1-Naphthol could be of concern because it is

more toxic than carbaryl to certain species, including freshwater fish (Tilak et al. 1981,

Brena et al. 2005).

Carbaryl is broken down by various pathways in the environment. Hydrolysis by

bacteria and aqueous photolysis are the most important processes for breakdown in water

(Brena et al. 2005). Carbaryl has a half-life of about four days under most environmental

conditions (Miller 1993). Other environmental factors can affect the rate of carbaryl

breakdown in the environment (e.g., increased sunlight and UV-B radiation, pH,

temperature, natural organic matter, nitrate) (Miller and Chin 2002), therefore the

duration of carbaryl in the environment is highly dependent upon environmental

conditions (Relyea 2003). For example, the pH of natural wetlands is generally 5.0-9.0;

at pH 9.0, the half-life of carbaryl is approximately 0.1 day, whereas it could take almost

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4 yrs for carbaryl to break down at pH 6.0 (Relyea 2003). The duration of exposure to

carbaryl is thus highly variable (Relyea 2003).

Direct Effects

Time-to-Death Assays

Time-to-death assays are toxicological tests that examine mortality at a single

concentration across time; thus measuring the rate of mortality as a result of chemical

exposure (Bridges and Semlitsch 2000). Bridges and Semlitsch (2000) measured time-to-

death in tadpoles under laboratory conditions to examine carbaryl sensitivities among

nine ranid species, within a single species (i.e. among populations), and within

populations (i.e. among families) of a single species.

Significant differences in time to death among the nine ranid species were

observed. Rana aurora was the most tolerant species followed by R. pretiosa, R.

palustris, R. sphenocephala, R blairi, R. clamitans, R. boylii, R. areolata, and R. sylvatica

was the most sensitive to carbaryl. It is interesting to note that two of the three western

U.S. species that have experienced the greatest declines, R. pretiosa and R. aurora were

the most tolerant of carbaryl; while the third, R. boylii, had average tolerance. Bridges

and Semlitsch (2000) attributed this observation to these species becoming locally

adapted to harsh environmental conditions (i.e. pesticide exposure).

Time to death also varied significantly among R. sphenocephala populations

indicating significant variation in carbaryl tolerance within populations throughout the

species’ range (Bridges and Semlitsch 2002). The differences in sensitivity were

21

suggested to be a result of “lingering effects of a past environmental selective pressure of

unknown intensity,” (Bridges and Semlitsch 2002).

Semlitsch et al. (2000) conducted time-to-death assays on Hyla versicolor

tadpoles under laboratory conditions to measure carbaryl tolerance of tadpoles from

different families and correlated data with mean performance of each family reared in

field enclosures (i.e. enclosed natural ponds). Significant variation in carbaryl tolerance

was observed among tadpoles within a single population of the gray treefrog, Hyla

versicolor. Semlitsch et al. (2000) believed that the intrapopulation differences in

carbaryl tolerance, although affected by genetic variation in body size, more likely

developed from environmental maternal contributions (e.g., egg size and yolk quality),

which could be important in amphibian population dynamics. Further, data indicated that

genotypes tolerant to chemical contamination may be less fit in other ways as a result of

natural selection. This study demonstrated that understanding environmental and genetic

variation in chemical tolerance is critical to identifying species vulnerability to

contaminants and thus understanding species declines (Semlitsch et al. 2000).

LC50 Toxicity Assays

Relyea and Mills (2001) demonstrated that results of LC50 tests may be

significantly overestimated. For example, Bridges (1997) estimated that the

concentration necessary to kill 50% of the population in two days (LC502-d) for Hyla

versicolor to be 12.9 mg/L. The LC504-d for Hyla versicolor, determined by Zaga et al.

(1998), was 2.5 mg/L, which is consistent with LC50 estimates for other larval anurans

(Marian et al. 1983; Marchal-Segault 1976). However, Relyea and Mills (2001) found

22

that carbaryl concentrations of only 0.05 mg/L and 0.09 mg/L caused up to 97% mortality

of H. versicolor tadpoles over 10 d (i.e. half of the population was exterminated in about

7 d). Therefore, lower concentrations of carbaryl may cause mass mortality of

amphibians, only over a longer period of time. Thus, LC50 tests should be conducted

using low and environmentally relevant concentrations, and testing periods should be

extended by at least a few days, to obtain more exact estimates of acute toxicity (Relyea

and Mills 2001).

Relyea (2003) used five concentrations of carbaryl (0.03, 0.3, 1.6, 3.2, and 6.5

mg/L) to determine LC50 estimates for six anuran species of tadpoles in the absence and

presence of predator cues (see Indirect Effects below). The LC50 estimates for the direct

effects of carbaryl (chronic) exposure on Rana sylvatica, Rana pipiens, Rana clamitans,

Rana catesbeiana, Bufo americanus, and Hyla versicolor were 1.2 mg/L, 2.2 mg/L, 2.6

mg/L, 2.3 mg/L, 3.4 mg/L, and 2.5 mg/L, respectively. Previous LC50 values, obtained

over shorter exposure durations (3-4 d), ranged from 2.5 to 18 mg/L (Marchal-Segault

1976, Marian et al. 1983, Bridges 1997, Zaga et al. 1998) whereas LC50 values ranged

from 1.2 to 3.4mg/L in this study (Relyea 2003). The LC50 estimates of anurans

obtained in this study may be more appropriate for natural wetlands where the pH is low

and carbaryl breakdown is slower (see Concentration and Breakdown in the

Environment, above) (Relyea 2003).

Sublethal Effects

Carbaryl is found in field concentrations (�4.8mg/L, Norris et al. 1983) generally

lower than those that are directly lethal to amphibian larvae (Boone et al. 2001).

23

Therefore it is important to study the impact of the pesticide at low, sublethal

concentrations (Bridges, 1999). The sublethal effects of a pesticide are the effects of

chemical exposure that is not toxic enough to cause direct mortality. Sublethal levels of

direct carbaryl exposure have the potential to alter population dynamics because they can

cause amphibian species to have a shorter larval period and increased rates of

metamorphosis, associated with smaller size (Bridges 1997). Carbaryl may directly affect

metamorphosis by activating corticotropin-releasing horomone, a stress horomone that

initiates premature metamorphosis in tadpoles in drying and stressed conditions (Denver

1995). Earlier metamorphosis is correlated with lower overwinter survival, lower

lifetime reproduction, and suppressed immune function (i.e. increased vulnerability to

disease) (Smith 1987; Carey et al. 1999). Further, larval performance is highly correlated

with adult health, and, thus, survival (Wilbur 1997). Therefore, low levels of carbaryl

could cause a population to decline slowly over time (Bridges and Semlitsch 2000).

Sublethal effects may be more valuable in assessing sensitivity to contaminants than

lethal effects (Little et al. 1990) because the sublethal effects may be more detrimental to

amphibian populations than effects of direct mortality (Peacor and Werner 2001).

Rohr et al. (2003) conducted static renewal laboratory tests that exposed

streamside salamander (Ambystoma barbouri) embryos to 0.5, 5, and 50 µg/L carbaryl

for 37 d. The experiment was developed to represent the direct effects of very low

sublethal levels of carbaryl believed to persist in aquatic ecosystems for significant

periods of time (Rohr et al. 2003, Gibbs et al. 1984). Carbaryl, at 50 µg/L, produced

asymmetrical larvae with missing limbs and digits, and caused significant mortality in

salamander larvae; thus A. barbouri could be highly sensitive to contaminants (Rohr et al.

24

2003). Also, there was greater mortality in larvae than embryos, which Rohr et al. (2003)

consider to be a result of either bioaccumulation (i.e. accumulation of chemicals over

time) or greater larval susceptibility to chemical exposure (i.e., jelly coating of eggs could

offer protection to embryos). Carbaryl also caused larvae to spend more time in refuge

and decreased activity. Lethargic larvae were least likely to display standard responses to

hunger. The sublethal effects of carbaryl exposure (e.g., deformities, lethargy, increased

use of refuge) could cause less effective foraging, increased predation, suppressed

immune response, and ultimately decreased survival in streamside salamanders (Rohr et

al. 2003).

Bridges (1997) exposed tadpoles to sublethal levels of carbaryl to monitor the

direct effects of the insecticide exposure on general activity and swimming performance.

Activity and swimming performance was recorded for each individual tadpole prior to

chemical exposure and at 24, 48, 72, and 96 h for the duration of exposure. Then tadpoles

were placed in fresh water and monitored at 24 and 48 h post-exposure.

Carbaryl caused nearly 90% reduction in activity at 3.5 mg/L, a relevant

environmental concentration. Tadpole activity decreased thirty minutes after exposure

and did not change during subsequent observations; therefore the effects of carbaryl are

demonstrated within a very short time after introduction into the aquatic environment

(Bridges 1997). At 48 h post-exposure, tadpole activity recovered for subjects exposed to

3.5mg/L, but significant recovery was not observed at higher concentrations. Bridges

(1997) concluded that acute exposure to sublethal levels of carbaryl produces a reduction

in both the activity and swimming performance in R. blairi tadpoles. The effects

25

observed in this study showed that carbaryl may strongly impact critical tadpole life

history functions and indirectly affect adult fitness and survival (Bridges 1997).

Bridges and Semlitsch (2000) monitored spontaneous swimming activity of ranid

tadpoles exposed to a 2.5 mg/L. Decreased activity levels were observed in all nine ranid

species exposed to carbaryl. Activity levels also differed among eight R. sphenocephala

populations exposed to carbaryl, indicating differential sensitivities throughout the

species range (Bridges and Semlitsch 2000). Variation was not observed among families

of R. sphenocephala (Bridges and Semlitsch 2000).

Saura-Mas et al. (2002) exposed Hyla versicolor eggs, and tadpoles to 3.5 mg/L

carbaryl to test for sublethal effects on survival, mass, and time to metamorphosis. The

anurans were exposed to the chemical in the laboratory for 48 hours; then, the subjects

either remained in the laboratory or were transferred to field enclosures (e.g. natural

ponds enclosed to hold tadpoles and exclude competitors and predators).

The study found no direct effect of chemical exposure on tadpoles. However,

animals reared in the laboratory had longer larval periods, larger mass at metamorphosis,

and better survival than those reared in the field, which were likely effects of

competition, predator cues (e.g., of predator presence outside of enclosures) and

fluctuating abiotic factors (e.g., temperature, pH, dissolved oxygen) under field

conditions. Thus, the effects observed in laboratory studies may be different than the

effects of the toxicant in the field (see Field Studies and Multiple Stressors below).

Further, the dominant effects of the insecticide on amphibians in the aquatic environment

are believed to be indirect effects (Boone and Semlitsch 2001, 2002). The pesticide

exposure was not necessarily inconsequential, but may have affected endpoints not

26

considered in this study (Saura-Mas et al. 2002). While single-factor studies are

important, they do not necessarily reveal the consequences that pesticides have in the

environment (Mills and Semlitsch 2002).

Indirect Effects

Laboratory Studies

The effects of carbaryl interact with other environmental factors or alter the

biological community thereby having indirect effects on a species (Mills and Semlitsch

2004). Bridges (1999) exposed H. versicolor tadpoles to two sublethal levels of carbaryl

(1.25 and 2.50 mg/L) expected to occur in the environment post-application, to determine

effects on tadpole activity and predator avoidance behavior. Bridges (1999) observed

that chemical exposure caused tadpoles to reduce use of refugia in the presence predator

(i.e., adult Notophthalmus viridiscens) chemical cues, thus increasing risk of predation.

Tadpoles exposed to carbaryl also spent more time in refugia when a predator cues were

not present, thereby limiting their food resources. Carbaryl exposure also caused an

overall decrease in activity of tadpoles, and produced longer larval periods and smaller

size at metamorphosis. The effects observed in this study could diminish adult fitness,

performance, and survival (Wilbur 1997, Smith 1987).

Boone and Bridges (1999) tested the effects of temperature on carbaryl potency

using Rana clamitans tadpoles. Tadpole survival was negatively correlated with the

interaction of temperature, chemical concentration, and time. Tadpoles exposed to high

concentrations of carbaryl had lower survival rates at 27ºC than at 17ºC and 22ºC, but at

low chemical concentrations, tadpoles survival was unaffected by temperature (i.e.

27

temperature treatment alone was not lethal). At high temperatures, a lower concentration

of carbaryl is needed to induce mortality. Because many amphibians breed and develop

in waters that commonly reach 27ºC or greater, carbaryl may be more potent in the

environment than expected (Boone and Bridges 1999). Carbaryl is absorbed by the body

more rapidly at high temperatures, but metabolism also increases with temperature,

therefore the chemical may be broken down and excreted more rapidly as well (Boone

and Bridges 1999). Furthermore, although carbaryl toxicity increases with temperature,

the chemical also breaks down more quickly in the aquatic environment at higher

temperatures, thus amphibians may be exposed to a more toxic insecticide but for a

shorter period of time. This study demonstrated the importance of testing chemicals at a

range of temperatures.

Relyea and Mills (2001) exposed Hyla versicolor tadpoles to sublethal levels of

carbaryl and chemical cues of a predator, Ambystoma maculatum, enclosed within a cage.

Carbaryl concentrations (in the absence of predators) of only 0.09 and 0.05 mg/L caused

high mortality within one week, most mortality occurring after 5 days. At 0.09 mg/L,

survival declined to about 8% by day 8, regardless of predator treatment. At 0.05 mg/L,

carbaryl caused survival to decline to 40% in the absence of predators, and to 3% in the

presence of predators. At concentrations of 0.07, 0.14, 0.27, and 0.54 mg/L, predator

cues caused carbaryl to be four times more lethal; survivorship averaged 32% across all

treatments. The interaction of carbaryl and predator cues induced mortality, significantly

reduced tadpole activity, and reduced tadpole growth by 50% compared to controls. So,

very low concentrations of carbaryl can drastically affect amphibian behavior, growth,

and survival (Relyea and Mills 2001).

28

Relyea (2003) then exposed anuran species to five concentrations of carbaryl and

predator cues (i.e. caged adult newt, Notophthalmus viridescens) to test for synergistic

interactions (i.e. indirect effects) and estimate LC50 values in the presence and absence

(see Direct Effects above) of predator cues. LC50 values in the presence of predator cues

for Rana pipiens, Rana clamitans, Rana catesbeiana, Bufo americanus, and Hyla

versicolor were 2.2 mg/L, 1.1 mg/L, 1.0 mg/L, 3.4 mg/L, and 2.5 mg/L, respectively.

Synergistic interactions between carbaryl and predator cues were not detected in

Rana sylvatica and Hyla versicolor. In fact, wood frogs (Rana sylvatica) experienced

significant mortality from predator cues in the absence of carbaryl; thus the LC50

estimate was not determined for indirect effects on this species (Relyea 2003). For Rana

pipiens and Bufo americanus, synergistic interactions were apparent early in the

experiments at high concentrations, but synergies disappeared because carbaryl alone

eventually killed all the tadpoles. Highly synergistic interactions persisted in R.

clamitans and R. catesbeiana—predator cues caused carbaryl to be up to 8 and 46 times

more deadly in R. clamitans and R. catesbeiana, respectively (Relyea 2003).

A noteworthy observation is that synergies were detected in two Missouri

populations of H. versicolor using chemical cues of Ambystoma maculatum (Relyea and

Mills 2001), but were not found in the Pennsylvania population, using N. viridescens cues

(Relyea 2003). Therefore, either different predators cause different synergistic effects or

different populations have different susceptibilities to the interactions of carbaryl and

predator cues (Relyea 2003). Because there are significant potential interactions between

chemical contaminants and predatory stress, further investigation must evaluate the

synergistic relationships of the stressors.

29

Field Studies

Laboratory tests are valuable in assessing the direct toxicity of chemical to an

amphibian species; however the results of these tests may not accurately represent the

effects of chemical contamination in the environment (Relyea 2005). For instance,

laboratory studies have shown negative direct effects of carbaryl on anuran

metamorphosis and survival (e.g., reduced survival and smaller size at metamorphosis)

(Bridges 1997, 1999). In contrast, mesocosm studies, which test the effects of the

contaminant on experimental aquatic communities, using a natural food web, have shown

positive indirect effects on anuran metamorphosis and survival (e.g., increased survival

and larger size at metamorphosis) resulting from increased periphyton (i.e. food)

abundance and decreased predation (i.e. reduced predator survival) (Mills & Semlitsch

2004; Boone & Semlitsch 2001, 2002, 2003). In these studies, the indirect benefits to the

anurans that result from changes in the aquatic community prevailed over the direct

consequences of chemical toxicity to the species. Mesocosm studies that replicate

contamination events on a natural aquatic ecosystem by utilizing natural biotic factors

such as plankton (i.e. food), natural predators and competitors, as well as abiotic factors

such as temperature, pond drying (i.e. hydroperiod), and pH, are therefore credited more

suitable to represent the effects of chemical exposure on a species in its natural

environment (Bridges and Semlitsch 2000, Mills 2002, Boone and Bridges-Britton 2006).

Notopthalmus viridescens larvae, although not directly affected by a concentration

of carbaryl, suffer mass mortality by carbaryl contamination of the aquatic environment

(Mills 2002; Mills and Semlitsch 2003). When the plankton that they depended upon for

30

nourishment were exterminated by the chemical, the salamanders died of starvation

(Mills 2002; Mills and Semlitsch 2003). Similar effects have been observed for

Ambystoma maculatum growth and survival (Boone and James 2003; Boone et al. 2007).

Thus, it is essential to understand the effects of the chemical on other organisms that exist

within the amphibian’s natural environment. “When toxicity studies are embedded in the

nexus of interactions that compose natural food webs, we can arrive at very different

interpretations due to the prevalence of both direct and indirect effects” (Relyea 2005).

It is essential to include natural factors such as competition, predation, and pond

hydroperiod (Boone and James 2003) as they are important in community processes

(Semlitsch et al. 1996), and understanding the effects of a chemical is impossible without

considering the complex environments that are also altered with chemical exposure

(Boone and James 2003). Therefore, not only must scientists consider appropriate

chemical concentrations, but it is also imperative to consider natural biological conditions

when determining the impacts of insecticides on amphibians (Relyea 2005).

Multiple Stressors

Environmental stressors such as increased UV-radiation and pathogens can

interact with pesticides to have synergistic effects on amphibian species by increasing

chemical toxicity or by increasing the vulnerability of the victim to other factors; thus, it

is necessary for studies to examine the role of multiple stressors in amphibian

communities (Boone and Bridges-Britton 2006). A single stressor is unlikely to act alone

in an aquatic ecosystem; consequently the effects of the stressor may be dependent upon

its interactions with other environmental factors (Boone et al. 2007).

31

Combinations of contaminants in the presence of other stressors have been shown

to have additive (Boone and James 2003) and synergistic (Boone et al. 2005, Relyea

2004, Hayes et al. 2003) interactions, although multiple, sublethal chemical stressors may

be no more damaging than the presence of a single chemical (Relyea 2005, Boone and

Bridges-Britton 2006, Puglis and Boone 2007). Combinations of multiple chemical

stressors in the environment could likely increase amphibian susceptibility to other

environmental factors, therefore causing population declines, (Carey et al. 1999, Boone et

al. 2007) but the combination of stressors may not produce effects that are predictable by

the effects observed by a single stressor (Boone et al. 2008).

Amphibians exposed to one agrichemical are likely to be exposed to more than

one. Crop maintenance, in farming practices as well as general lawn and garden care,

generally requires the use of fertilizers in addition to herbicides and insecticides. Boone

et al. (2005) found that carbaryl in the presence of ammonium nitrate, directly affected

Rana clamitans by creating greater metabolic demands, leaving less energy for feeding or

food assimilation thereby, reducing growth and development. Boone and Bridges-Britton

(2006), however, reported no additive effects associated with the combination of carbaryl

and ammonium nitrate exposure on Hyla versicolor.

Boone et al. (2007) tested the interactions of (2.5 mg/L) carbaryl and (10mg/L)

ammonium nitrate fertilizer on an experimental aquatic community comprised of Bufo

americanus tadpoles, Rana sphenocephala tadpoles, and Ambystoma maculatum

salamander larvae. Overwintered Rana catesbeiana bullfrogs served as competitors with

the tadpoles and bluegill Lepomis macrochirus were top predators. Boone et al. (2007)

did not observe negative synergistic interactions between the chemicals. In fact, the

32

insecticide and the fertilizer increased periphyton (i.e. food) abundance which enhanced

anuran survival and increased mass at metamorphosis. Rana sphenocephala exposed to

either bullfrogs or bluegill had decreased time to metamorphosis, while tadpoles exposed

to both bullfrogs and bluegill increased time to metamorphosis.

Ambystoma maculatum were affected by carbaryl treatment and the interaction of

carbaryl and bluegill which reduced cladoceran (i.e. food) abundance thereby reducing

salamander mass, increasing time to metamorphose, and causing mortality to the

salamanders in the aquatic community (Boone et al. 2007). Carbaryl exposure likely

compromises growth, reproduction, and survival in A. maculatum in the environment.

Boone et al. (2007) suggests that A. maculatum may be most able to tolerate additional

biotic stress in the absence of chemicals.

Boone et al. (2001) and Boone and Bridges (2003) exposed Rana clamitans

tadpoles to sublethal concentrations of carbaryl to test the effects of multiple exposures to

the insecticide and timing of exposure. The number of times tadpoles were exposed to

carbaryl significantly increased the rate of metamorphosis, and decreased metamorph size

(Boone et al. 2001, Boone and Bridges 2003). Carbaryl caused mortality to tadpoles

exposed to carbaryl one time late in development, while tadpole survival was greater to

tadpoles exposed three times: one time during early, mid- and late development (Boone

and Bridges 2003). Boone (2008) asserted that pesticides with the same mode of action

(e.g., cholinesterase inhibitors) are less likely to produce additive effects from multiple

exposures in comparison to chemicals with different modes of action.

The indirect effects of carbaryl exposure at concentrations expected to occur in

the environment may have positive effects on anurans due to increased food resources

33

(Boone et al. 2008); however the effects of an additional stressor in the aquatic

community could reduce anuran diversity. For instance, Boone and Semlitsch (2003)

found that carbaryl enhanced the size and development of bullfrogs (Rana catesbeiana),

while the chemical induced predator (Notophtalmus viridescens, Lepomis macrochirus,

and Orconectes sp.) mortality presumably due to reduction in zooplankton food

resources. Bullfrogs, especially introduced populations, can be damaging to other

amphibian species by acting as competitors, predators, or disease vectors (Boone et al.

2008). Carbaryl exposure could allow bullfrogs to invade new communities by reducing

or eliminating predators, which could have negative consequences for other anurans in

the community (Boone and Semlitsch 2003, Boone et al. 2008). These data could help

explain bullfrog success in “highly managed areas,” (Boone and Semlitsch 2003, Boone

et al. 2008).

Interactive effects of UV-B radiation and chemical stressors are essential to

toxicity studies because UV-radiation alone can threaten amphibian survival and in the

presence of a pesticide, it may increase chemical toxicity. Zaga et al. (1998) tested the

interactive effects of UV radiation and carbaryl treatment on embryos and tadpoles of

Hyla versicolor and Xenopus laevis. Synergism between UV radiation and carbaryl

appeared to occur by photomodification of the carbaryl molecule in water (Zaga et al.

1998). The study found that carbaryl absorbs UV-A as well as UV-B radiation, and UV-

A also photoactivates the chemical, but to a lesser extent than with UV-B.

While 100% mortality occurred in X. laevis embryos exposed to carbaryl and high

UV-B radiation simultaneously, mortality was not observed in X. laevis embryos

previously exposed to carbaryl and subsequently exposed to UV-B radiation; therefore

34

the photochemical transformation of carbaryl likely produced the synergistic effects

observed in all experiments (Zaga et al. 1998). A ten-fold increase in toxicity resulted

from photoactiviation of carbaryl by approximately 1.5% ambient UV-B (Blaustein et al.

2003). Increased swimming activity, followed by decreased locomotor activities, and

developmental abnormalities, such as lateral flexure of the tail, were also observed in X.

laevis and H. versicolor tadpoles exposed to UV-B radiation and carbaryl concentrations

of 0.86, 1.24, and 1.76 mg/L (Zaga et al. 1998). Hence, the acceptable level of carbaryl

in the environment, using H. versicolor as the model species, may be much below 1.24

mg/L.

The interaction between pesticides and pathogens is also important to the study of

amphibian population declines. These chemicals may inhibit immune defenses and

increase susceptibility to disease (Davidson et al. 2007). Puglis and Boone (2007)

exposed Rana catesbeiana embryos and tadpoles to the pathogen, Saprolegnia ferax, and

sublethal levels of carbaryl and ammonium nitrate under laboratory conditions.

Significant interactive effects between the pathogen, the fertilizer, and the insecticide

were not observed, although similar concentrations of carbaryl were found to interact

with other amphibian species (Puglis and Boone 2007).

Davidson et al. (2007) exposed post-metamorphic juvenile Rana boylii to the

chytid fungus (Batrachochytrium dendrobatidis) and sublethal levels of carbaryl.

Although carbaryl exposure inhibited peptide defenses in Rana boylii juveniles, no

significant direct effects on survival or growth were apparent from the interaction

between chytid and carbaryl exposure (Davidson et al. 2007). Davidson et al. (2007)

reported that the results could be due to either species or localized resistance to the strain

35

of chytid used in the study and that R. boylii may be more susceptible to different strains

of chytid than the one in this study. Further studies are needed to examine the interactive

effects of pathogens and carbaryl.

Amphibians are continuously responding to multiple abiotic (e.g., temperature,

UV-radiation, pond drying) and biotic (e.g., food resources, predation, competition,

density, pathogens) stressors. The complex interactions among these factors thus

determine amphibian survival. In order for conclusions to be made concerning the

relative impacts of agrichemicals on amphibian populations, further research must be

employed to examine the interactive effects of agrichemicals and environmental stressors

on aquatic communities. Stressors relating to current global change and amphibian

declines (e.g., UV-B radiation, temperature change, pathogens contributing to EIDs) are

critical to further studies.

Conclusion

The increased use of chemical pesticides is believed to contribute to the decline in

amphibian biodiversity. It is necessary to determine how the agrichemicals are impacting

amphibian populations in order to slow the rate of amphibian declines (i.e. reduce human

impacts from farming practices) and prevent irreversible damage to ecosystems. If

pesticides are contributing to declines, it is not likely due to direct lethal effects, but

sublethal effects and interactions with other stressors (Davidson et al. 2007). Single

factor studies may be insufficient to determine the widespread phenomena of amphibian

declines (Boone and James 2003). Because amphibian declines are generally believed to

be caused by multiple stressors, it is logical to conclude that better understanding of the

36

declines will therefore be found through the examination of the interactive effects of

those stressors.

In combination with multiple stressors, toxicology studies must also include

sublethal concentrations of the pesticides, which could interfere with population

dynamics and produce indirect effects caused by change in the aquatic community.

Study of these effects are essential to pesticide toxicity research and should be utilized in

chemical regulation. Also, in order to gain understanding about local adaptation and

persistence of species in contaminated environments, it is necessary to determine the

presence of genetic variation for tolerance in populations (Semlitsch et al. 2000).

Toxicology studies that assess the susceptibility to contaminants and other stressors in the

environment among different species and populations are necessary for monitoring

populations and could also aid in pesticide regulation. Additional studies on the additive

effects of multiple chemical exposures, timing (i.e. life stage) and frequency of

exposures, and chemical mixtures are necessary as well to link toxicology studies with

amphibian exposure in the environment.

Long-term toxicology studies and monitoring programs are critical to future

research. It is important to study multiple chemical exposures in aquatic communities for

multiple generations to gain understanding of population dynamics. Long-term studies

examining the shifts in community structure after the disappearance of key species could

also provide beneficial data. Finally, further studies to assess changes in species habitat

use as a result of contamination in the environment are needed. Storfer (2003) stressed

the use of monitoring programs that are statistically sensitive to changes in amphibian

populations. By determining the distribution and abundance of amphibian populations,

37

researchers will be more capable of assessing the threat and the extent of population

declines. Conservation efforts should also be made to monitor the landscape to determine

the extent and abundance of suitable habitats (Storfer 2003). Finally, molecular genetic

techniques should be employed to monitor demographics and identify significant events

such as population fragmentation and hybridization (Storfer 2003). Together, these

techniques should help to recognize major issues contributing to amphibian declines,

identify which species are most threatened, and aid in slowing the rate of amphibian

species loss. Furthermore, by employing the techniques described above, researchers

may be able to recognize patterns, make generalizations, and create a more standard

mechanized approach to understanding amphibian declines.

38

Acknowledgements

First, I would like to thank Frank Nelson for giving me the opportunity to work in

conservation and especially for introducing me to the topic of amphibian declines. It has

been a pleasure working for you. I would also like to thank Dr. Indi Braden. I cannot

express my gratitude to you for seeing my potential and always believing in me. Thank

you so much for all your encouragement, for displaying a personal interest in my

education, and for helping me to achieve my goals. I sincerely appreciate the help of Dr.

Stephen Overmann, Dr. Matthew Fasnacht, Dr. Michael Aide, and Dr. William

Eddleman. Your helpful suggestions and cooperation were essential to this project.

Thank you to my husband, Casey Mills, for never once doubting my capabilities, for

pushing me to work harder, and never letting me give up. I thank my daughter, Kaya, for

being so patient and tolerant of me. From your wisdom I have learned so much, and I am

anxious to learn more as you grow older. You are as luminous and as precious as the sun.

I want to thank my parents, Paul Barber and Brenda Barber, for being proud of me and

reminding me that I don’t have to be the best at everything, though you know I’ll never

give up trying. I cannot express my gratitude for the opportunities that you have

provided me and for your constant support over the years. I love you both and I am

fortunate to have parents like you. To my sister, Erin Asher, thanks for always listening.

I appreciate your compassion and your advice. Finally, I would like to thank Jack and

Robin Mills for their support. I am honored to be a part of your family.

39

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