Ecological Applications, 20(8), 2010, pp. 2263–2272 Ó 2010 by the Ecological Society of America
Interactions of environmental stressors impact survival and development of parasitized larval amphibians J. KOPRIVNIKAR1 Department of Biology, Brandon University, John R. Brodie Science Centre, 270 18th Street, Brandon, Manitoba R7A 6A9 Canada
Abstract. Infected hosts are exposed to many environmental stressors that must be taken into account in order to determine the importance of disease, as various combinations can interact in unpredictable ways. Here, northern leopard frog (Rana pipiens) tadpoles, a species in decline, were exposed to stressors singly or in combination. Stressors included infection by Echinostoma trivolvis (a trematode parasite), exposure to predator chemical cues (larval dragonflies), and exposure to varying concentrations of the herbicide atrazine. Parasitism decreased survival only in combination with exposure to 3 lg/L atrazine, with a negative interaction observed for mass as well. Similarly, a negative interaction of parasitism and predation on survival occurred. However, atrazine exposure alone negatively affected the survival, mass, and developmental stage of tadpoles. These results indicate that certain stressor combinations are particularly deleterious for young parasitized tadpoles. Notably, very common low-intensity parasite infection can be particularly harmful in certain situations. Such negative impacts on larval amphibians in certain scenarios may contribute to ongoing amphibian population declines, emphasizing that the combination of environmental stressors must be considered when evaluating the general role of disease in species extinctions. Key words: amphibian; atrazine; disease; Echinostoma trivolvis; environment; northern leopard frog; parasite; predator; Rana pipiens; stress.
INTRODUCTION Global species extinctions continue to occur at a high rate, driven by various environmental changes such as habitat loss, pollution, and introduced species (see Sala et al. 2000, Baillie et al. 2004 for reviews). However, infectious disease has not traditionally been regarded as a significant driver of species extinctions and has been found to be significantly less likely than other drivers to act in isolation (Smith et al. 2006), even though mathematical models and epidemiological theory suggest a strong impact of infectious diseases on host populations (e.g., Anderson and May 1992). In isolation, the effects of parasitism on hosts are likely to be important for population regulation, but interactions with other driving factors may contribute to local and global extinctions (Smith et al. 2009). As such, the importance of disease for conservation efforts is likely best thought of in the context of disease ecology, i.e., considering hosts, pathogens, and the environment as systems of interacting agents. One potentially very important interaction for hosts is that between pathogens and environmental pollutants. The majority of studies have focused on effects of pollutants for host susceptibility to disease, and an Manuscript received 26 August 2099; revised 14 January 2010; accepted 19 January 2010. Corresponding Editor: J. Van Buskirk. 1 E-mail:
[email protected]
increasing number of studies show that common environmental pollutants may impair the immune system of a wide range of animal taxa (Selgrade 2007). However, we must also consider how already infected hosts fare in the presence of additional stressors such as pollutants. In recent years, there have been an increasing number of papers indicating that parasitism and pollution can interact with each other (see Lafferty 1997, Sures 2008 for reviews). The presence of contaminants appears to be particularly detrimental for infected vs. uninfected individuals in many studies (e.g., Hecker and Karbe 2005, Coors and De Meester 2008). The importance of considering multiple stressors in relation to species declines is particularly relevant for amphibians, with over 100 species suspected to have gone extinct in recent decades, and many others considered threatened or endangered (Stuart et al. 2004, Collins and Crump 2009). While many factors have been implicated in this phenomenon (e.g., contaminants, habitat loss), amphibian population declines are likely due to multiple causes that may interact (Blaustein and Kiesecker 2002), and natural populations are often exposed to more than one stressor concurrently such that they may interact synergistically (Sih et al. 2004). As such, the context in which amphibians are exposed to environmental stressors must be taken into account, as various combinations of such stressors could be the underlying cause of observed declines (Linder et al. 2003, Stuart et al. 2004).
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With respect to amphibian disease, the emergence of the fungal pathogen Batrachochytrium dendrobatidis has had devastating effects on populations, and is considered one of the greatest threats to global amphibian diversity (Skerratt et al. 2007). Another prominent pathogen is the parasitic trematode Ribeiroa ondatrae, a platyhelminth that causes elevated mortality and severe malformations in larval amphibians (e.g., Johnson et al. 1999, Johnson and Hartson 2009). Parasites are in fact a ubiquitous component of natural ecosystems, and individual frogs may support many trematode species and several thousand individual parasites (see Sutherland 2005); however, the impact of parasitic infection on amphibian populations may be strongly context dependent (Kiesecker and Skelly 2001). In light of continuing worldwide amphibian declines, it is essential that we determine the role of pathogens (Daszak et al. 2003), necessitating the examination of potential interactions of parasitism with additional stressors and their importance for amphibian populations. Here I examined the interaction of trematode (Echinostoma trivolvis) infection, exposure to the herbicide atrazine, and exposure to predator (larval Anax dragonflies) chemical cues on the mass and developmental stage of northern leopard frog (Rana pipiens) tadpoles. Notably, northern leopard frogs represent a species in decline throughout much of its range (Rorabaugh 2005), and the co-occurrence of trematode infection and exposure to agrochemicals may be common for this species (e.g., Rohr et al. 2008). Atrazine is a frequent contaminant of surface and subterranean waters (Solomon et al. 1996) and is commonly detected in agricultural areas supporting amphibian populations (e.g., Koprivnikar et al. 2006a, Rohr et al. 2008). Various studies have reported a variety of effects on amphibians, from decreased growth and survivorship to accelerated or delayed development (see Rohr and McCoy 2009 for a review). Atrazine has also been shown to increase larval amphibian susceptibility to trematode infection (Kiesecker 2002), but it is not known how infected hosts respond to atrazine exposure in comparison to uninfected individuals. However, it is possible that interactive effects could occur, given that the presence of an additional stressor (predators) causes sublethal concentrations of atrazine and other pesticides to become lethal to larval frogs (e.g., Relyea and Mills 2001, Relyea 2004, LaFiandra et al. 2008). Aquatic prey often respond to predators via chemical cues released by the predator and/or prey during or after the predation event (see Chivers and Smith 1998 for a review). Predator presence affects stress hormones and immune function in vertebrates (Boonstra et al. 1998, Horak et al. 2006), and reduces energy storage levels (Stoks et al. 2005). In combination with other stressors, predation may be an additional stressor on amphibian physiology that results in negative synergistic effects (Relyea and Mills 2001). As such, the combination of
parasite infection, atrazine exposure, and exposure to predator cues may interact in ways that cannot be predicted from single-factor experiments but could have significant implications for host condition and survival, which may translate into population- or species-level effects. MATERIALS
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METHODS
Experimental design The experiment had a two-by-two-by-three complete factorial design with 32 replicates (individual tadpoles). The three factors were atrazine exposure (0, 3, and 300 lg/L), predation cue (present, absent), and trematode parasite exposure ( yes, no), which resulted in 12 treatments in total (excluding the diluted solvent control). The individual treatment containers were randomly assigned positions along two adjacent laboratory benches in the same room. Parasite exposure Tadpoles were exposed to a free-living infective stage of the parasitic trematode Echinostoma trivolvis. Following asexual reproduction within a first intermediate host (the aquatic gastropod Planorbella/Helisoma trivolvis here), free-living infective larval stages (cercariae) of E. trivolvis emerge and seek out a suitable second intermediate host (including larval amphibians) wherein they develop into cysts (metacercariae) within the kidneys after migrating up the cloaca. After infected tadpoles/frogs are ingested by the definitive host (various birds or mammals for E. trivolvis), the metacercariae develop into adult worms and reproduce sexually (Olsen 1974). Northern leopard frog eggs were obtained from the Carolina Biological Supply Company (Burlington, North Carolina, USA) in early October 2008 and raised in six 20-L aquaria filled with dechlorinated tap water. Six batches of eggs were received, but it is unknown whether these came from the same clutch. Approximately 125 tadpoles were kept in each aquarium and were fed boiled spinach and rabbit chow ad libitum for roughly six weeks until they reached Gosner developmental stage 25–26 (Gosner 1960). At this time, ;75 tadpoles were haphazardly chosen from each aquarium and pooled together into a single bucket, resulting in a pool of over 450 tadpoles. Tadpoles were then individually transferred to 140-mL plastic cups containing 100 mL of dechlorinated tap water. Only those tadpoles without hind limb buds obvious to the naked eye, defined as less than Gosner developmental stage 27 here, were retained for the experiment (N ¼ 450). Half of the cups contained 15 cercariae obtained from naturally infected P. trivolvis collected in Santa Clara County, California, USA. Cercariae were obtained as detailed in Koprivnikar et al. (2008) and were identified using a key by Schell (1985) and by the fact that they used P. trivolvis as a host. This number of cercariae was chosen as mortality is highly likely when leopard frog
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tadpoles are exposed to as few as 20 cercariae while in Gosner developmental stage 25 (Schotthoefer et al. 2003). The small volume of water was used to preclude any anti-parasite behaviour by tadpoles (Koprivnikar et al. 2006b). Tadpoles were kept in the cups overnight before being transferred into their experimental containers the next day. Note that 34 tadpoles died during the parasite exposure period (21 in cups with cercariae, 13 cups without cercariae), leaving 416 tadpoles available for the remainder of the experiment. Pesticide exposure Technical grade atrazine (Pestanal analytic standard: 99.2% pure) was obtained from Sigma Aldrich (St. Louis, Missouri, USA) and a stock solution with a concentration of 1 mg/mL was prepared using 95% ethanol (see Griggs and Belden 2008). I then diluted this first stock solution by adding 1 mL of it to 999 mL of dechlorinated water to obtain a second stock solution with a concentration of 1000 lg/L atrazine. The lower atrazine concentration (3 lg/L atrazine) was then attained by adding 45 mL of the second stock solution to 14 L and 955 mL of dechlorinated tap water in a plastic carboy. The higher atrazine concentration (300 lg/L atrazine) was achieved by adding 4.5 mL of the first stock solution to 14 L and 995.5 mL of dechlorinated tap water in another plastic carboy. I chose the lower concentration to reflect the maximum allowable atrazine level in drinking water as determined by the U.S. Environmental Protection Agency (Aspelin 1997). The higher atrazine concentration reflects a probable field scenario, given that a recent metaanalysis of atrazine effects considered concentrations near or below historical environmental concentrations of ;500 lg/L to be ecologically relevant for the purposes of the study (Rohr and McCoy 2009). The control consisted of dechlorinated tap water. I also included a diluted solvent control of 100 lg/L EtOH for comparison to the control, given that the atrazine was dissolved using ethanol. Of the 416 tadpoles in the experiment, 32 were assigned to the solvent control. After parasite exposure as detailed above, tadpoles were transferred into individual 2-L plastic containers holding either 1 L of dechlorinated tap water, 1 L of solvent control solution, 1 L of 3 lg/L atrazine solution, or 1 L of 300 lg/L atrazine solution. As previous research has demonstrated that there is only a minor loss of atrazine in freshwater aquaria over a one-week period (Rohr et al. 2004), water changes took place every seven days. Water changes maintained treatments, given that refills were consistent with the original treatment/control solution. Predator cue exposure Larval Anax dragonflies were obtained from Ward’s Natural Science (Rochester, New York, USA). For the predation events, three pairs of dragonfly larvae were kept in separate 3-L containers holding either 2 L of
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dechlorinated tap water, 2 L of 3 lg/L atrazine solution, or 2 L of 300 lg/L atrazine solution, along with small rocks to use as refugia. Larval dragonflies have welldocumented effects on tadpole growth and development, particularly via chemical cues present after prey ingestion (see Benard 2004, Relyea 2007 for reviews). To obtain predator chemical cues, I placed two tadpoles into each larval container and left them overnight. I transferred the Anax larvae into new containers containing dechlorinated tap water the next morning in the event that they might be affected by atrazine exposure and also rotated the dragonfly larvae through the three different atrazine concentrations during successive predation events. Note that all tadpoles were consumed during each predation event, and that all prey tadpoles were approximately the same size (;0.5 g; small to encourage predation). Half of the tadpoles in the 2-L containers then received 40 mL of strained predator water appropriate to their treatment (e.g., those in 3 lg/L atrazine received predator cue in 3 lg/L atrazine), while the other half received 40 mL of water also appropriate to their treatment, resulting in final holding volume of 1.04 L for each individual tadpole. Addition of predator cue/sham occurred on the same day as water changes, i.e., every seven days. A recent study has shown that predator chemical cues from larval dragonflies degrade in 2–4 days (Peacor 2006). Tadpole maintenance and examination Each tadpole received a standard-sized (1 cm length) pellet of rabbit chow after each water change. The room in which all the experimental tadpoles were housed was kept at a constant temperature of 208C, with a light : dark cycle of 14:10. Tadpoles were monitored daily, with any dead tadpoles recorded and removed, until the termination of the experiment, which occurred when the first tadpole undergoing tail resorption was observed (approximately Gosner stage 43). It should be noted that this was one of only two individuals to exceed Gosner stage 39 at the termination point. At this time, tadpoles were euthanized in a buffered solution of MS-222, weighed, and preserved in 95% EtOH. Determination of the Gosner developmental stage (Gosner 1960) took place afterward, along with dissections to enumerate the number of cysts in tadpoles exposed to cercariae. Four tadpoles from each of the treatments without parasite exposure were also dissected to confirm that none had kidney metacercariae. Statistical analysis All analyses were conducted using SPSS 17.0 (SPSS 2009). Tadpole mass was log-transformed to meet the assumption of normality. A one-way ANOVA was used to test for differences between the solvent control and control with respect to mass and developmental stage, while a chi-square test using the crosstabs procedure was used to check for a difference in mortality. As no significant differences were found (see Results), the
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survival. Two sets of analyses were performed for the mass and stage data: One excluded tadpoles that were exposed to cercariae but subsequently found to be uninfected, while the other included such individuals. This was done because tadpoles that actually harbored cysts at the end of the experiment were far more likely to show effects of parasitism. Even though larval amphibians exhibit behavioral responses in the presence of cercariae (e.g., Koprivnikar et al. 2006b), it is unknown whether such brief exposure to cercariae without host penetration actually constitutes a physiological stressor that could potentially interact with other stressors. The exclusion of uninfected tadpoles in one set of analyses thus allows for the specific examination of stressor interactions with parasite infection vs. parasite exposure. As such, 204 tadpoles were included when exposed but uninfected individuals were excluded, while 230 were included when data for all surviving individuals were used. Mass and developmental stage were analyzed with a multivariate analysis of variance (MANOVA) using the general linear model procedure to determine whether E. trivolvis exposure/infection, atrazine exposure, predator cue exposure, or interactions among these factors had an overall effect on tadpole mass and developmental stage, followed by univariate tests. A MANOVA was used as these measures were unlikely to be independent of one another. RESULTS Mortality
FIG. 1. Mortality of tadpoles of the northern leopard frog (Rana pipiens) under different treatments of (a) atrazine exposure and (b) exposure to a trematode parasite, Echinostoma trivolvis.
solvent control data was pooled with the control for further analyses. The effects of varying atrazine exposure on mortality were analyzed using the crosstabs procedure and included all 416 tadpoles. The interactive effects of the stressors for mortality were examined using a factorial logistic regression via the forward conditional method with trematode exposure, exposure to 0 or 3 lg/L atrazine, and predator cue exposure as covariates and status as dead or alive at the end of the experiment as the dependent factor. This was done because very few tadpoles exposed to 300 lg/L atrazine survived until the end of the experiment; thus, it was not appropriate to attempt to detect interactions of exposure to 300 lg/L atrazine with additional stressors given that there was essentially no variation in mortality due to additional exposure to parasites or predator cue. Mortality in 300 lg/L atrazine ranged from 95% to 100%. With respect to effects on mass and developmental stage, tadpoles exposed to 300 lg/L atrazine were also excluded from these analyses, given their very low
There was no significant difference in mortality between the control and solvent control (Pearson v2 ¼ 0.988, df ¼ 1, P ¼ 0.320) as survival was 78.1% in the control and 87.5% in the solvent control. Results of the ANOVA also indicated no significant difference with respect to mass (F1,51 ¼ 1.584, P ¼ 0.214) and developmental stage (F1,51 ¼ 1.597, P ¼ 0.212). The mean cyst intensity for infected tadpoles was 7.6 6 0.25 (all data shown are means 6 SE), with a range of 6–14 metacercariae. A significant difference among the three atrazine concentrations was observed for tadpole survival (Pearson v2 ¼ 205.543, df ¼ 2, P , 0.0001; Fig. 1). The model (v2 ¼ 24.155, df ¼ 6, P , 0.0001) resulting from the factorial logistic regression examining separate and interactive stressor effects included exposure to 3 lg/L atrazine (Wald statistic ¼ 6.384, P ¼ 0.012) and exposure to parasites (Wald statistic ¼ 6.235, P ¼ 0.013), as well as significant interactions between parasite and predator cue exposure (Wald statistic ¼ 7.187, P ¼ 0.007), and between exposure to parasites and 3 lg/L atrazine (Wald statistic ¼ 8.634, P ¼ 0.003). Mortality was lower in parasite-exposed tadpoles but higher in tadpoles exposed to 3 lg/L atrazine; however, parasite-exposed tadpoles experienced increased mortality when additionally exposed to 3 lg/L atrazine or predation cues while those not exposed to parasites did not (Fig. 2).
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Mass and developmental stage Results from the multivariate tests including parasiteexposed but uninfected tadpoles showed significant overall effects of parasite exposure (Wilks’ lambda ¼ 0.969, df ¼ 2, 221, P ¼ 0.030) and exposure to 3 lg/L atrazine (Wilks’ lambda ¼ 0.808, df ¼ 2, 221, P , 0.0001). No significant effects were observed for predation alone (Wilks’ lambda ¼ 0.999, df ¼ 2, 221, P ¼ 0.869), nor were there significant interactions between parasite and predator cue exposure (Wilks’ lambda ¼ 0.998, df ¼ 2, 221, P ¼ 0.995), exposure to parasite and to 3 lg/L atrazine (Wilks’ lambda ¼ 0.983, df ¼ 2, 221, P ¼ 0.155), exposure to predator cue and 3 lg/L atrazine (Wilks’ lambda¼ 0.985, df¼ 2, 221, P ¼ 0.185), or among all three factors (Wilks’ lambda ¼ 1.0, df ¼ 2, 221, P ¼ 0.957). Results from the univariate ANOVAs indicated that parasite exposure had an effect by negatively impacting developmental stage (F1, 222 ¼ 4.683, P ¼ 0.032), with parasite-exposed tadpoles having a mean developmental stage of 33.9 6 0.2,while those not exposed to cercariae had a mean of 34.5 6 0.2. Exposure to 3 lg/L atrazine had a significant overall effect by decreasing both mass (F1, 222 ¼ 42.034, P , 0.0001) and developmental stage (F1, 221 ¼ 46.949, P , 0.0001). Tadpoles exposed to 3 lg/L atrazine had a mean log-mass of 0.011 6 0.017 compared to 0.129 6 0.014 for unexposed tadpoles, and those exposed to 3 lg/L atrazine had a mean developmental stage of 33.3 6 0.2 compared to 35.2 6 0.2 for unexposed tadpoles (see Appendix A for all treatment means). Results from the multivariate tests that excluded parasite-exposed but uninfected tadpoles showed significant overall effects of parasite infection (Wilks’ lambda ¼ 0.942, df ¼ 2, 195, P ¼ 0.003) and exposure to 3 lg/L atrazine (Wilks’ lambda ¼ 26.659, df ¼ 2, 195, P , 0.0001). In addition, significant interactions were seen between parasite infection and exposure to 3 lg/L atrazine (Wilks’ lambda ¼ 0.969, df ¼ 2, 195, P ¼ 0.046), and between exposure to predator cue and 3 lg/L atrazine (Wilks’ lambda ¼ 0.958, df ¼ 2, 195, P ¼ 0.016). No significant effects were observed for predation alone (Wilks’ lambda ¼ 0.998, df ¼ 2, 195, P ¼ 0.789), nor were there significant interactions between parasite infection and predator cue exposure (Wilks’ lambda ¼ 0.999, df ¼ 2, 195, P ¼ 0.936) or among all three factors (Wilks’ lambda ¼ 0.992, df ¼ 2, 195, P ¼ 0.464). Results from the univariate ANOVAs indicated that parasite infection had an effect by negatively impacting developmental stage (F1, 196 ¼ 11.455, P ¼ 0.001), with infected tadpoles having a mean developmental stage of 33.5 6 0.2, while those not infected had a mean of 34.5 6 0.2. Exposure to 3 lg/L atrazine had a significant overall effect by decreasing both mass (F1, 196 ¼ 43.348, P , 0.0001) and developmental stage (F1, 196 ¼ 45.923, P , 0.0001). Tadpoles exposed to 3 lg/L atrazine had a mean log-mass of 0.039 6 0.019 compared to 0.122 6 0.015 for unexposed tadpoles, and those exposed to 3 lg/L atrazine had a mean developmental stage of 33.0 6 0.2 compared to 35.0 6 0.2 for unexposed tadpoles. The
FIG. 2. Tadpole mortality due to (a) parasite and predator cue (larval Anax dragonflies) exposure, and (b) parasite and 3 lg/L atrazine exposure. Notations of and þ indicate absence and presence, respectively, of exposure to factor.
significant interaction between parasite infection and exposure to 3 lg/L atrazine occurred because the mass of infected and uninfected tadpoles did not differ in 0 lg/L atrazine, but the decrease in mass resulting from exposure to 3 lg/L atrazine was stronger for infected tadpoles such that their mean mass was significantly less than that of uninfected tadpoles (Fig. 3). The significant interaction between predator cue and 3 lg/L atrazine exposure occurred because predator cue-exposed tadpoles were significantly less developed than those not exposed to predator cues in the absence of atrazine, but this was reversed in the presence of atrazine, even though atrazine exposure negatively impacted developmental stage for both (Fig. 3). DISCUSSION The results of the current study emphasize the need to consider context when determining the importance of pathogens for host populations. While exposure to parasitism had little overall effect on larval amphibians here, it is clear that this stressor in combination with others can have interactive effects and thus such phenomena should be a critical consideration when considering the impacts of pathogens and other stressors on threatened and endangered species.
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FIG. 3. (a) Mean mass (untransformed) of parasite-infected (þ) and uninfected () tadpoles in the presence of 3 lg/L atrazine (dot-dashed line) or 0 lg/L atrazine (solid line). (b) Mean Gosner developmental stage of tadpoles exposed (þ) or unexposed () to a predator cue in the presence of 3 lg/L atrazine (dot-dashed line) or 0 lg/L atrazine (solid line). Differentiation of the hind-limb buds begins at Gosner stage 31, and forelimbs emerge at Gosner stage 42. Error bars represent 6SE. Parasite-exposed but uninfected tadpoles were not included for either measure.
While parasite exposure/infection did not affect mortality, parasite-exposed/infected tadpoles were less developed than those not exposed/infected, which is not unexpected given previous experiments with E. trivolvis and leopard frog tadpoles (e.g., Schotthoefer et al. 2003). Experimentally infected tadpoles can suffer edema, intensity-dependent mortality, inhibited growth, and an inflammatory response (Martin and Conn 1990, Fried et al. 1997). However, the negative effect on development seen here is somewhat surprising given the relatively low mean intensity of infection and the finding that moderate infection by this trematode has no effects on physiology and fitness-related traits of larval pickerel frogs (Orlofske et al. 2009). One possible explanation is interspecific variation among larval amphibians in parasite effects, as observed for the trematode R. ondatrae (Johnson and Hartson 2009), thus this should be further explored. It is interesting to note that results
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from analyses including or excluding parasite-exposed but uninfected tadpoles appear to differ with respect to interactions with other stressors. This indicates that while there may be a cost of any possible resistance to infection or simply being exposed to cercariae, there is a higher cost of actually harboring cysts. As such, infection by, rather than simple exposure to, parasite infectious stages appears to be more important in the context of synergistic interactions. However, this phenomenon should be further explored given that tadpoles exhibit a ‘‘fear response’’ simply in the presence of cercariae as exhibited by hyperactivity induced by the mere detection of cercarial chemical and/or vibrational cues (Rohr et al. 2009). The finding that exposure to both 3 and 300 lg/L atrazine increased mortality indicates a greater sensitivity to this concentration than previously reported, as exposure to 3 lg/L atrazine does not usually significantly decrease survival (see Rohr and McCoy 2009 for a review). There are, however, exceptions to this trend, such as the report by Storrs and Kiesecker (2004) that exposure to 3 lg/L atrazine results in higher mortality than exposure to 30 or 300 lg/L atrazine for four larval amphibian species (northern leopard frogs not included). The LC50 value (defined as the concentration required to kill half the members of a tested population after a specified test duration; also known as the median lethal dose) for late-stage northern leopard frog tadpoles has been reported to be 14 500 lg/L atrazine; however, this was based on a 96-hour exposure period (Howe et al. 1998) and the effects of short-term vs. long-term exposure can be very different (Storrs and Kiesecker 2004). Larval pickerel frogs (Rana palustris) exposed to a single dose of 117 lg/L atrazine over four weeks exhibited significantly increased mortality; however, this was not observed for green frog (Rana clamitans) larvae (Rohr et al. 2008). As there is substantial variation among amphibian species in their response to other pesticides as well (e.g., Relyea 2009), further studies involving long-term exposure of leopard frog tadpoles to atrazine are needed in order to firmly establish the sensitivity of this species in comparison to others. Reduced mass and developmental stage resulted from exposure to 3 lg/L atrazine here; however, it is unlikely that smaller tadpoles were selected against via decreased survivorship in atrazine, given that a significant decrease in mass was still observed. A recent meta-analysis of atrazine effects on size at metamorphosis shows a clear dose-dependent response, as atrazine exposure was consistently associated with size reductions at ecologically relevant concentrations (Rohr and McCoy 2009). While a detailed examination of the proposed mechanisms behind atrazine’s detrimental effects is beyond the scope of this paper, pesticides can primarily have negative effects on life history by reducing food intake (Ribeiro et al. 2001), or decreasing conversion efficiency, likely due to reallocation of energy to other processes like detoxification (Congdon et al. 2001). It has also
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been proposed that atrazine may affect corticoid and thyroid hormones (e.g., Hayes 2000, Brodeur et al. 2009), which are critical to the timing of amphibian metamorphosis. The meta-analysis by Rohr and McCoy (2009) also found a number of studies reporting significant effects of atrazine on metamorphic timing, but the direction of effect was not consistent across studies, and some had either clear or possible nonmonotonic dose responses. On its own, predation had no effect on mortality, mass, or developmental stage. A recent review found that chemical cues emitted by predators rarely induce a smaller size at metamorphosis or a shorter time to metamorphosis despite theoretical predictions (Relyea 2007); thus, the results here are generally consistent with those previously reported. In addition to the main effects, the impacts of predator cue and parasite exposure were also often dependent on the presence of additional stressors. Predation cues alone had no significant effect, but individuals exposed to predation cues were more developed than those not exposed to predator cues in 3 lg/L atrazine, while the opposite pattern was observed in the absence of atrazine. No significant interaction of exposure to predation cue and atrazine for mortality was observed. This is inconsistent with results reported by Lafiandra et al. (2008), but a lower atrazine concentration was used in the present study. Numerous other studies have also found negative synergistic effects on amphibian mortality arising from simultaneous exposure to predation stress and sublethal concentrations of pesticides (e.g., Relyea and Mills 2001, Relyea 2003). As such, the nature of the interaction observed here indicates the possibility that perception of the predator chemical cues may have been altered in the presence of atrazine; however, reports indicate that olfactory ability in amphibians is not impacted by atrazine exposure (see Rohr and McCoy 2009 for a review). Alternatively, rapid breakdown or alteration of the predator cue may have occurred in the presence of atrazine. The most likely explanation for the observed effect on developmental stage is that tadpoles may have accelerated development to escape what may have been perceived as a very poor environment (Wilbur and Collins 1973). The lack of an interaction of predator cue and atrazine exposure for mortality may be due to the fact that predation is likely to have been a noncontinuous stressor here compared to the design of other studies (e.g., Relyea and Mills 2001). Similarly, while predation cues alone had no overall impact on tadpole mortality, infected individuals experienced higher mortality when additionally exposed to this stressor, representing the first report of such an interaction. Here predation cues likely simply represented an additional physiological stressor. As such, the combination of parasitism and predation can not only increase host predation risk (e.g., through altered host
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behavior; see Moore 2002 for a review), but also increase host mortality in the absence of predation. With respect to the interaction between parasite and atrazine exposure, infected tadpoles exhibited decreased survival when additionally exposed to 3 lg/L atrazine, while those that were uninfected did not. In fact, increased mortality due to exposure to this atrazine concentration seems to be driven by effects on infected individuals. In addition, there was no difference in mean mass between infected and uninfected tadpoles in the absence of atrazine, but infected individuals showed a greater decrease in mass in response to atrazine exposure than those uninfected. This indicates that low levels of parasitism are unlikely to strongly affect such earlystage tadpoles, but that the addition of a contaminant stressor can result in particularly detrimental effects. In particular, while it has been demonstrated that exposure to 3 lg/L atrazine can increase tadpole susceptibility to parasitism (Kiesecker 2002), the current results indicate that exposure of already infected individuals to this concentration of atrazine early in their development also has negative consequences. This is in keeping with the increasing number of reports showing that pollution enhances negative effects of parasites or that parasites interfere with hosts’ protection mechanisms against pollutants, thereby increasing harmful effects of toxic substances (see Sures 2008 for a review). Interactions among pathogens and contaminants have been noted for other aquatic vertebrates (e.g., Jacobson et al. 2003, Marcogliese et al. 2005, Sures 2008) as well as amphibians. For example, the highest levels of stress biomarkers in bullfrogs (Rana catesbeiana) were found in individuals from highly agricultural areas that were severely infected with a particular trematode species (Marcogliese et al. 2009). However, Budischak et al. (2009) detected no effect of E. trivolvis infection on tadpole susceptibility to the insecticide malathion, but noted that it is important to investigate other pesticides, pathogens, and amphibian hosts before dismissing this type of interaction. Similarly, while the insecticide carbaryl significantly reduced tadpole survival and hatching rate, there was no interaction with a pathogenic fungus (Puglis and Boone 2007). As such, it is highly likely that interactive effects on amphibians are both pathogen- and stressor-dependent and require further exploration. The population-level consequences of pathogen– stressor interactions with respect to amphibian declines are not easy to determine as parasitism alone may have significant effects. Beasley et al. (2005) reported that as much as 50% of kidney tissue was occupied by echinostome trematode cysts in northern cricket frogs (Acris crepitans), and suggested that this might be related to declines in cricket frog recruitment in wetlands with high echinostome abundance. As several studies have found a negative relationship between amphibian abundance and proximity to agricultural land (e.g., Davidson et al. 2001, Davidson 2004), it is a distinct
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possibility that a combination of pesticide and pathogen exposure could be particularly detrimental and play a role in population declines if infected hosts experience this additional stressor, given the impact of each factor on its own. Agricultural ponds are becoming an important breeding habitat (Knutson et al. 2004), and many ‘‘natural’’ aquatic systems are also contaminated with at least one pesticide (Gilliom et al. 2007). As parasite infection in amphibians is often higher in agricultural settings in which pesticides are also commonly detected (e.g., Koprivnikar et al. 2006a, Rohr et al. 2008), infected individuals may face particularly adverse consequences in response to continued low-level atrazine exposure early in development. Consideration of the context in which larval amphibians experience environmental stressors is critical to conservation efforts. The combination of stressors must be included as we attempt to elucidate the major drivers behind amphibian population declines. The results reported here support the idea that amphibian population declines are likely due to interacting multiple causes (Blaustein and Kiesecker 2002), and that the major contribution of pesticides is likely due to sublethal effects and interactions with other stressors (Carey and Bryant 1995). Consequently, amphibian populations/ species experiencing such situations, such as those whose early larval development coincides with atrazine application and a high likelihood of parasite infection, may be of particular concern. While lethal effects, such as the increased mortality resulting from stressors alone or in combination seen here, can clearly impact amphibian populations, traitmediated effects of stressors are also critical for population regulation and should not be overlooked. In particular, smaller size at metamorphosis can have profound effects on juvenile frog survival and reproductive potential. Smaller individuals are more susceptible to predators (Wilbur and Collins 1973, Skelly 1994, Relyea 2003), and reduced size at metamorphosis also delays reproductive maturity and decreases fecundity (Wilbur and Collins 1973, Berven and Gill 1983). Negative interactions between pathogens and environmentally relevant contaminant concentrations could thus indirectly regulate populations by affecting mass as seen here in addition to directly affecting the survival of amphibian larvae. Given the results of this study and others examining host–pathogen–environment interactions, the overall importance of disease as a driver of species declines and extinctions must take into account how infected hosts fare when exposed to additional stressors in addition to effects on host susceptibility to disease. As environmental changes such as habitat loss and contamination continue to occur, the resulting implications for host disease should be explored in a manner that incorporates a realistic consideration of the situations that threatened species are likely to experience. For example, along with exposure to predation and pollut-
ants, altered species diversity could also have profound consequences for host disease dynamics (e.g., Keesing et al. 2006), as well as the introduction of nonnative species (Kelly et al. 2009, Johnson and Thieltges 2010). The potential for pathogens to significantly impact endangered and threatened species cannot be properly evaluated without a disease ecology framework that includes interactions among hosts, their pathogens, and their environment; thus, further studies incorporating such dynamics are required to elucidate the role of disease as a driver of species declines and extinctions. ACKNOWLEDGMENTS I thank F. Shaheen, D. Y. Lim, C. Fu, M. G. Tien, T. T. Le, and S. H. Brack at the University of the Pacific for assistance in maintaining, dissecting, and staging tadpoles, as well as J. R. Rohr and M. D. Boone for comments on an earlier version of this manuscript. This study was supported by an Eberhardt Research Fellowship to J. Koprivnikar. LITERATURE CITED Anderson, R. M., and R. M. May. 1992. Infectious diseases of humans, dynamics and control. Oxford University Press, New York, New York, USA. Aspelin, A. L. 1997. Pesticide industry sales and usage. Environmental Protection Agency, Washington, D.C., USA. Baillie, J. E. M., C. Hilton-Taylor, and S. N. Stuart, editors. 2004. 2004 IUCN red list of threatened species. A global species assessment. IUCN, Cambridge, UK. Beasley, V. R., S. A. Faeh, B. Wikoff, J. Eisold, D. Nichols, R. Cole, A. M. Schotthoefer, C. Staehle, M. Greenwell, and L. E. Brown. 2005. Risk factors and the decline of the northern cricket frog, Acris crepitans: evidence for involvement of herbicides, parasitism, and habitat modifications. Pages 75–87 in M. J. Lannoo, editor. Amphibian declines: the conservation status of United States species. University of California Press, Berkeley, California, USA. Benard, M. F. 2004. Predator-induced phenotypic plasticity in organisms with complex life cycles. Annual Review of Ecology and Systematics 35:651–673. Berven, K. A., and D. E. Gill. 1983. Interpreting geographic variation in life-history traits. American Zoologist 23:85–97. Blaustein, A. R., and J. M. Kiesecker. 2002. Complexity in conservation: lessons from the global decline of amphibian populations. Ecology Letters 5:597–608. Boonstra, R., D. Hik, G. R. Singleton, and A. Tinnikov. 1998. The impact of predator-induced stress on the snowshoe hare cycle. Ecological Monographs 68:371–394. Brodeur, J. C., G. Svartz, C. S. Perez-Colla, D. J. G. Marinob, and J. Herkovits. 2009. Comparative susceptibility to atrazine of three developmental stages of Rhinellaarenarum and influence on metamorphosis: non-monotonous acceleration of the time to climax and delayed tail resorption. Aquatic Toxicology 91:161–170. Budischak, S. A., L. K. Belden, and W. A. Hopkins. 2009. Relative toxicity of malathion to trematode-infected and noninfected Rana palustris tadpoles. Archives of Environmental Contamination and Toxicology 56:123–128. Carey, C., and C. J. Bryant. 1995. Possible interrelations among environmental toxicants, amphibian development, and decline of amphibian populations. Environmental Health Perspectives 103:13–17. Chivers, D. P., and R. J. F. Smith. 1998. Chemical alarm signalling in aquatic predator-prey systems: a review and prospectus. Ecoscience 5:338–352. Collins, J. P., and M. L. Crump. 2009. Extinction in our times: global amphibian decline. Oxford University Press, Oxford, UK.
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APPENDIX Treatment means of mass and development stage for tadpoles exposed to trematode parasites, predator cues, and 3 lg/L atrazine (Ecological Archives A020-085-A1).
