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Effects of chlorothalonil on development and growth of amphibian embryos and larvae
Environmental Pollution 181 (2013) 329e334
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Environmental Pollution journal homepage: www.elsevier.com/locate/envpol
Short communication
Effects of chlorothalonil on development and growth of amphibian embryos and larvae Shuangying Yu a, Mike R. Wages a, George P. Cobb b, Jonathan D. Maul a, * a b
Department of Environmental Toxicology, The Institute of Environmental and Human Health, Texas Tech University, Lubbock, TX 79416, USA Department of Environmental Science, Baylor University, Waco, TX 76798, USA
a r t i c l e i n f o
a b s t r a c t
Article history: Received 3 January 2013 Received in revised form 15 May 2013 Accepted 15 June 2013
Chlorothalonil is a broad spectrum fungicide widely used in agricultural and urban environments, yet little is known regarding its effects on amphibians. We examined effects of chlorothalonil on growth, malformations, and mortality in embryos and larvae of Xenopus laevis and Spea multiplicata, and assessed variation in sensitivity among aquatic organisms using a species sensitivity distribution (SSD). Chlorothalonil induced gut malformations in X. laevis embryos and inhibited growth. Tail degeneration was observed in larvae of both species and reduced tail length to total length ratios occurred at environmentally relevant concentrations (5.9 and 11.0 mg/L). The mechanism of tail degeneration is unclear, but alteration in the expression of genes involved in tail resorption is a hypothesized mechanism. Larval amphibians were more sensitive than invertebrates and fish. Based on our results and the range of reported environmental concentrations, chlorothalonil may pose a risk to larval amphibians in certain habitats and scenarios. Ó 2013 Elsevier Ltd. All rights reserved.
Keywords: African clawed frog Fungicide Malformation Mexican spadefoot toad Tail degeneration
1. Introduction Chlorothalonil (2,4,5,6-tetrachloroisophthalonitrile) is the most commonly used fungicide in the U.S. (USEPA, 2011) and is widely used in agriculture to protect crops (USEPA, 1999). Chlorothalonil may be introduced to aquatic habitats by direct application, spray drift, and runoff, which may pose a risk to aquatic organisms. Chlorothalonil concentrations in runoff ranged from 50 to130 mg/L after two days of rainfall (Potter et al., 2001) and can be as high as 500 mg/L during the first runoff event (Wilson et al., 2010). Average body residue in Pacific treefrog (Hyla regilla) tadpoles collected from the Kaweah River basin, CA ranged from 33.3 to 47.7 ng/g wet weight (Datta et al., 1998). Chlorothalonil is highly toxic to fish and invertebrates (USEPA, 1999). The 24e96 h LC50s ranged from 12 to 195 mg/L for freshwater invertebrates and 16 to 76 mg/L for freshwater fish (van Wezel and van Vlaardingen, 2004). Chlorothalonil is toxic to tunicate blood cells at 1 mM (Cima et al., 2008) and fish phagocytes at 250 mg/L (Baier-Anderson and Anderson, 1998). Chronic exposure at 2.0 mg/L can decrease hematocrit and cause gill damage in fish (Davies, 1987). However, studies that address chlorothalonil toxicity to amphibians are limited. A 48-h LC50 of 160 mg/L was reported for * Corresponding author. E-mail address: [email protected] (J.D. Maul). 0269-7491/$ e see front matter Ó 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.envpol.2013.06.017
Japanese common toad Bufo bufo japonicus tadpoles (Hashimoto and Nishiuchi, 1981). Recently, adverse effects on survival, liver tissue and immune cells, and corticosterone concentrations in anurans have been reported at chlorothalonil concentrations between 0.0164 and 164 mg/L (McMahon et al., 2011). Taken together, these studies indicate that chlorothalonil may have a significant impact on amphibians and toxicity data are needed. The objectives of this study were to: (1) examine effects of chlorothalonil on development and growth in two anuran species, African clawed frogs (Xenopus laevis) and Mexican spadefoot toads (Spea multiplicata), and (2) compare sensitivity of larval amphibians and other aquatic organisms to chlorothalonil. 2. Materials and methods 2.1. Experiment 1: embryo and larval Xenopus laevis 96-h acute toxicity tests Breeding, housing, and field collection methods for X. laevis and S. multiplicata have been described in the supplemental materials. Toxicity tests were conducted following standardized FETAX methods (ASTM, 2004) with slight modifications. Embryos were not dejellied to achieve environmental realism and because egg jelly may protect embryos from ambient contaminants (Edginton et al., 2007). Embryos were collected from three clutches, mixed, and examined for viability and developmental stage. Twenty NF stage 8e11 embryos (Nieuwkoop and Faber, 1975) were randomly assigned to each of 50 ml glass jars containing 40 ml FETAX solution spiked with chlorothalonil. Concentrations used were solvent control (acetone), 7.8, 13.0, 21.6, 36.0, and 60.0 mg/L (see Supplemental materials for rationale for choosing these concentrations). There were five replicates for each treatment.
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S. Yu et al. / Environmental Pollution 181 (2013) 329e334
In the larval experiment, NF stage 46 larvae from three clutches were mixed and 10 larvae were randomly placed in 1 L glass jars containing 900 ml FETAX solution. Treatments included solvent control, 10.2, 12.8, 16.0, 20.0, and 25.0 mg/L, and there were five replicates for each treatment. To compare growth and other morphological measurement before and after exposure, 18 larvae were randomly collected from the same clutches prior to the exposure. These were used to quantify initial size of larvae. Larvae were fed approximately an hour before water change with a diet described in Koss and Wakeford (2000). Experiment 1 was conducted under static renewal conditions with daily water change. All animals were maintained on a 12:12, light:dark cycle at temperatures ranging from 22 to 24 C. Water quality was measured every other day. Embryos and larvae were checked daily for mortality (i.e., no response to prodding). At the end of the exposure, larvae were euthanized and preserved in formalin. Each individual was examined for abnormalities under a microscope. Images of each larvae were taken and total length (head to tail) (TL), body length (BL), and snout-to-vent length (SVL) were measured using Image J (version 1.44P, Wayne Rasband, National Institutes of Health, USA). Larvae exposed to chlorothalonil showed reduced tail length; therefore, tail length was also examined and calculated from TL and BL (Tail ¼ TL BL). We used the tail to TL ratio to minimize the effect of growth rate on absolute tail length measurements and compared the ratio among treatments. Initial larval size measurements prior to the exposure were included in comparisons to demonstrate growth in controls during the experimental period. 2.2. Experiment 2: verification experiments for Xenopus laevis To verify the results of Experiment 1, partial toxicity tests with three replicates per treatment were conducted for embryos and larvae with the same procedures and conditions as previous experiments. However, embryos from only two clutches were used due to lack of breeding or low quality eggs in other clutches. Three clutches were used for the larval experiment. Thirty-one larvae were euthanized and preserved in formalin before the exposure and used to determine initial size measurements. Larvae were fed daily 2 mg of SeraÒ Micro per experimental unit. 2.3. Experiment 3: larval Spea multiplicata 96-h acute toxicity test Larvae (Gosner stage 25) from five clutches were mixed and were randomly assigned to each treatment. Five larvae were placed in 500 ml glass jars with 400 ml laboratory water and six replicates were used for each treatment. A 96-h static non-
renewal toxicity test was performed. Treatments included solvent control, 3.2, 5.4, 9.0, 15.0, and 25.0 mg/L. Fifteen larvae were collected for initial size measurements. Larvae were not fed during the exposure. Water quality was measured at the end of the exposure. Morphological measurements were similar to X. laevis. 2.4. Comparison of chlorothalonil toxicity to amphibians, fish, and invertebrates A species sensitivity distribution (SSD) curve was created for fish, invertebrates, and amphibians to compare sensitivity among taxa and evaluate the risk chlorothalonil poses to aquatic organisms. The SSD curve was fit with the log-probit distribution using the U.S. EPA Species Sensitivity Distribution Generator (SSD_Generator_V1.xlt). We only used 96-h LC50s to minimize the variation due to exposure duration. 2.5. Chemical and statistical analyses Chlorothalonil exposure concentrations were quantified using a gas chromatograph equipped with an electron capture detector. Measured chlorothalonil concentrations were reported throughout with correction for extraction efficiency. Data were analyzed using JMP Statistical Analysis Software (Ver 9.0.0, Statistical Analysis System Institute, Cary, NC, USA) and results were considered significant at a ¼ 0.05 (see Supplemental materials for more information on chemical and statistical analyses).
3. Results and discussion 3.1. Chemical analyses Measured water concentrations of chlorothalonil ranged from 58 to 142% of the nominal concentrations with an average of 96% (Table 1). In Experiment 3, the mean chlorothalonil concentrations were 84, 84, and 76% of the initial concentrations for 3.2, 9.0, and 15.0 mg/L (nominal) treatments, respectively, after 96 h (data provided in Supplemental materials). Water quality data were also provided in Supplemental Materials.
Table 1 Nominal and measured chlorothalonil concentrations (mg/L) at the beginning of exposures, percent mortality at each concentration, 96-h median lethal concentrations (LC50s, 95% confidence interval), and model parameters associated with LC50 estimates for Xenopus laevis and Spea multiplicata. Species
Stage
Nominal (mg/L)
Measured (mg/L)
Mortality (%)
LC50 (95% CI)
Slope
Intercept
X. laevisa
Embryo
Control 7.8 13.0 21.6 36.0 60.0 Control 10.2 12.8 16.0 20.0 25.0
0 8.8 12.8 19.3 36.4 60.7 0 5.9 8.3 14.4 17.0 23.8
0 8 3 6 16 99 10 29 52 82 98 100
42.4 (39.8e45.3)
0.13
5.36
8.2 (7.1e9.2)
0.31
2.57
Control 7.8 13.0 21.6 36.0 60.0 Control 10.2 12.8 16.0 20.0 25.0
0 6.1 9.5 18.8 26.8 60.3 0 8.6 11.1 13.4 16.3 22.5
0 5 2 23 75 100 3 10 0 27 86 100
22.9 (21.4e24.6)
0.25
5.68
14.4 (13.6e15.4)
0.56
8.02
Control 3.2 5.4 9.0 15.0 25.0
0 4.6 7.0 11.0 20.5 34.6
0 0 0 7 93 100
10.7 (9.7e12.2)
0.58
6.26
Larvae
X. laevisb
Embryo
Larvae
S. multiplicata
a b
Larvae
Experiment 1. Experiment 2, verification experiment.
S. Yu et al. / Environmental Pollution 181 (2013) 329e334
3.2. Effects of chlorothalonil on survival The LC50s (95% CI) for X. laevis embryos in Experiment 1 and Experiment 2 were 42.4 mg/L (39.8e45.3) and 22.9 mg/L (21.4e24.6), respectively (Table 1). For X. laevis larvae, the LC50 was 8.2 mg/L (7.1e9.2) in Experiment 1 and 14.4 mg/L (13.6e15.4) in Experiment 2. The LC50 for S. multiplicata larvae was 10.7 mg/L (9.7e12.2), similar to those for X. laevis larvae. Chlorothalonil was less toxic to X. laevis embryos compared to X. laevis larvae, indicating differential sensitivity among developmental stages. Our 96-h LC50s for larvae of both species were approximately 10e20 times lower than the 48-h LC50 (160 mg/L) reported by Hashimoto and Nishiuchi (1981) for B. bufo japonicus, suggesting large variation in sensitivity to chlorothalonil among amphibian species. It should be noted, however, that exposure time varied between our study and that of Hashimoto and Nishiuchi (1981). 3.3. Effects of chlorothalonil on development Chlorothalonil produced malformations in X. laevis embryos, including gut malformations, axial/tail malformations, and edema. Proportions of malformed embryos were significantly different among treatments in both Experiment 1 (ANOVA, F4,20 ¼ 15.5, P < 0.001) and Experiment 2 (F4,10 ¼ 24.7, P < 0.001) (Fig. 1).
100
A
%Total malformations %Gut malformatinons %Axial/tail malformations %Edema
%Malformation
80
60
** 40
20
331
Compared to controls (mean SE: 4.92 2.20% and 9.24 2.50% in Experiment 1 and 2, respectively), incidence of total malformations was significantly higher at 36.4 mg/L (42.56 5.02%) in Experiment 1 and at 18.8 mg/L (41.11 8.04%) and 26.8 mg/L (93.33 6.67%) in Experiment 2 (Dunnett’s test, all P < 0.05). Chlorothalonil at 26.8 and 36.4 mg/L produced significantly greater proportions of gut malformations and/or axial/tail malformations compared to controls (all P < 0.05) (Fig. 1). Most interestingly, X. laevis larvae exhibited tail shortening during chlorothalonil exposure (Fig. 2). Because of the high mortality at the two highest concentrations (i.e., 100% mortality or data from only one replicate), these two treatments were not included in statistical analyses. Tail to TL ratios were significantly different among treatments in Experiment 1 (KruskaleWallis test, P < 0.001), and tail to TL ratios for all chlorothalonil concentrations (0.56 0.006) were significantly lower compared to control (0.63 0.002) and initial (0.64 0.003) (all P < 0.001) (Fig. 3A). In Experiment 2, larvae from 11.1 (0.61 0.007) and 13.4 mg/L (0.56 0.006) had significantly lower tail to TL ratios than control (0.66 0.001) and initial (0.67 0.004) (all P < 0.01) (Fig. 3B). Similar to X. laevis, we observed tail shortening in S. multiplicata, and larvae at 11.0 (0.57 0.003) and 20.5 mg/L (0.37 0.063) had significantly lower tail/TL ratio compared to control (0.60 0.002) (all P < 0.05) (Fig. 3C). Tail degeneration caused by chlorothalonil in both species is the most important finding of this study. External Ca2þ has been reported to induce apoptosis and tail shortening in tadpoles (Menon et al., 2000) and acetochlor has been demonstrated to reduce tadpole body area and alter expression of T3 dependent genes (Crump et al., 2002). But to our knowledge, no study has reported that pesticides or other chemicals cause such remarkable tail shortening in early larval stages (i.e., NF stage 46 or Gosner stage 25). Both X. laevis and S. multiplicata larvae exposed to chlorothalonil had shorter tails compared to the initial group from the same clutches collected prior to the exposure. This difference suggests that the observed tail shortening is likely a degeneration process rather than growth inhibition. The mechanism of regressing tails is unknown but may be related to apoptosis of tail tissue similar to tail absorption during metamorphosis (Nakajima et al.,
0 Control
8.8
12.8
19.3
36.4
B
**
100
%Malformation
80
60
* *
40
20
0 Control
6.1
9.5
18.8
26.8
Concentration (µg/L) Fig. 1. Incidence of total, gut, and axial/tail malformations and edema in stage 8e11 Xenopus laevis embryos exposed to chlorothalonil for 96 h in Experiment 1 (A) and Experiment 2 (B). Concentrations on the x-axis are measured concentrations. Error bars are SE and asterisks indicate a significant difference compared to controls (P < 0.05).
Fig. 2. Tail shortening in representative stage 46 Xenopus laevis larvae exposed to different concentrations of chlorothalonil for 96 h in Experiment 1 (from top to bottom: control, 5.9, 8.3, and 14.4 mg/L).
0.70
0.70
A
B
0.65
0.65
0.60
0.60
Tail/TL ratio
Tail/TL ratio
* * *
0.55
*
0.50 0.45
*
0.55 0.50 0.45
0.40
0.40 Initial
Control
5.9
8.3
14.4
17
Initial
Control
8.6
11.1
13.4
16.3
Concentration (µg/L)
Concentration (µg/L)
0.65
C
0.60
*
Tail/TL ratio
0.55 0.50 0.45
*
0.40 0.35 0.30 Initial Control
4.6
7
11
20.5
Concentration (µg/L)
Fig. 3. Tail/total length (TL) ratio of stage 46 Xenopus laevis larvae exposed to chlorothalonil for 96 h in Experiment 1 (A) and Experiment 2 (B); and Gosner stage 25 Spea multiplicata larvae exposed to chlorothalonil for 96 h in Experiment 3 (C). The initial group represents morphological measurements on a haphazardly selected subset of tadpoles preserved immediately prior to the exposure. There was only one replicate remaining at 17.0 mg/L in Experiment 1 and 16.3 mg/L in Experiment 2. Concentrations on the x-axis are measured concentrations. Error bars are SE and asterisks indicate a significantly different tail/TL ratio compared to controls (P < 0.05). 4.0
4.0
B
3.8
3.8
3.6
3.6
SVL (mm)
SVL (mm)
A
* 3.4
*
3.4
3.2
3.2
3.0
3.0
*
Control
8.8
12.8
19.3
36.4
Control
Concentration (µg/L)
C
* *
*
18.8
26.8
3.5
D
5.0
SVL (mm)
SVL (mm)
*
4.0
9.5
Concentration (µg/L)
5.0
4.5
6.1
4.5
* *
4.0
3.5
Initial
Control
5.9
8.3
14.4
Concentration (µg/L)
17
Initial
Control
8.6
11.1
13.4
16.3
Concentration (µg/L)
Fig. 4. Snout-to-vent length (SVL) of stage 8e11 Xenopus laevis embryos in Experiment 1 (A) and Experiment 2 (B) and stage 46 Xenopus laevis larvae in Experiment 1 (C) and Experiment 2 (D). Both experiments were 96 h chlorothalonil exposures. Concentrations on the x-axis are measured concentrations. Error bars are SE and asterisks indicate a significantly different SVL compared to controls (P < 0.05).
S. Yu et al. / Environmental Pollution 181 (2013) 329e334
2005). It is possible that chlorothalonil may interfere with pathways that trigger tail resorption by altering expression of genes involved in tail cell death (Brown et al., 1996) (see Supplemental materials for additional discussion).
333
greatest sensitivity to chlorothalonil and that additional amphibian species may be needed to fully evaluate the risk this fungicide may pose to amphibian populations. 4. Conclusions
3.4. Effects of chlorothalonil on growth There was a significant difference in SVL among treatments for X. laevis embryos in both Experiment 1 (F4,20 ¼ 10.4, P < 0.001) (Fig. 4A) and Experiment 2 (F4,10 ¼ 29.9, P < 0.001) (Fig. 4B). Embryos from 36.4 mg/L (3.42 0.04 mm) in Experiment 1 and embryos from 18.8 mg/L (3.53 0.09 mm) and 26.8 mg/L (3.03 0.04 mm) in Experiment 2 were smaller than controls (3.64 0.04 and 3.77 0.04 mm for Experiment 1 and 2, respectively, all P < 0.001). Xenopus laevis larvae collected from the initial group and all chlorothalonil concentrations (the two highest concentrations not included) were significantly smaller (4.43 0.05 mm) than larvae from the control treatment (4.94 0.06 mm, all P < 0.001) in Experiment 1 (Fig. 4C). In Experiment 2, larvae from the initial group (4.22 0.09 mm) and 13.4 mg/L (3.87 0.08 mm) were significantly smaller compared to the control group (4.96 0.01 mm, all P < 0.001) (Fig. 4D). For S. multiplicata larvae, only larvae from the initial group had smaller SVL (3.26 0.09 mm) compared to the control (4.16 0.04 mm, P < 0.001) (see Supplemental materials for the figure and additional discussion). 3.5. Comparison of chlorothalonil toxicity to amphibians, fish, and invertebrates The 96-h LC50s for fish and invertebrates were obtained from the U.S. EPA OPP Pesticide Ecotoxicity Database and existing literature (Davies and White, 1985; Ernst et al., 1991; Montforts, 1999; Sherrard et al., 2002) (data provided in Supplemental materials). Spea multiplicata and X. laevis were more sensitive than fish and invertebrates that had available 96-h LC50 data (Fig. 5). The estimated 5% hazard concentration (HC5, 95% CI) was 6.28 mg/L (3.52e 11.20), and the HC95 was 178.55 mg/L (100.06e318.59). When the amphibian LC50s were compared among taxa of freshwater organisms, the LC50s of X. laevis and S. multiplicata larvae were near the lower range of LC50s for fish, indicating a greater sensitivity to chlorothalonil. Comparisons among all freshwater taxa suggests that larval amphibians may have the
Proportion of species affected
1.2 Predicted relationship 95% Confidence interval Amphibians Fish Invertebrates
1.0
0.8
Ceriodaphnia dubia Ictalurus punctatus Lepomis macrochirus Cyprinus carpio Oncorhynchus mykiss
0.6
Galaxia auratus Pimephales promelas
0.4
Galaxia truttaceus Galaxia maculatus Paratya australiensis Astacopsis gouldi Xenopus laevis Spea multiplicata
0.2
0.0
1
10
100
1000
96-h LC50 (µg/L) Fig. 5. Species sensitivity distribution (SSD) for chlorothalonil among freshwater fish, invertebrates, and data from two amphibian species, Xenopus laevis and Spea multiplicata from the present study. All data used in the SSD are 96-h median lethal concentrations (i.e., LC50s).
Our results suggest that chlorothalonil is toxic to X. laevis and S. multiplicata, and X. laevis larvae were more sensitive to chlorothalonil than embryos. Our study reveals for the first time that chlorothalonil disrupts tail development and causes tail degeneration. Other sublethal effects include inhibition of growth and malformations. Based on acute toxicity data, amphibians appear to be more sensitive to chlorothalonil and may be at greater risk compared to other aquatic organisms. Further investigation focused on molecular and cellular pathways are needed to reveal the mechanism of tail shortening observed in the present study. Acknowledgments S Yu received funding from the Paul Whitfield Horn Fellowship provided by the University Women’s Club of Texas Tech University. Appendix A. Supplementary data Supplementary data related to this article can be found at http:// dx.doi.org/10.1016/j.envpol.2013.06.017. References American Society for Testing and Materials, 2004. Standard Guide for Conducting the Frog Embryo Teratogenesis Assay-Xenopus (FETAX). ASTM E1439e98 (2004). Baier-Anderson, C., Anderson, R.S., 1998. Evaluation of the immunotoxicity of chlorothalonil to striped bass phagocytes following in vitro exposure. Environmental Toxicology and Chemistry 17, 1546e1551. Brown, D.D., Wang, Z., Furlow, J.D., Kanamori, A., Schwartzman, R.A., Remo, B.F., Pinder, A., 1996. The thyroid hormone-induced tail resorption program during Xenopus laevis metamorphosis. Proceedings of the National Academy of Sciences 93, 1924e1929. Cima, F., Bragadin, M., Ballarin, L., 2008. Toxic effects of new antifouling compounds on tunicate haemocytes I. Sea-Nine 211 and chlorothalonil. Aquatic Toxicology 86, 299e312. Crump, D., Werry, K., Veldhoen, N., Van Aggelen, G., Helbing, C.C., 2002. Exposure to the herbicide acetochlor alters thyroid hormone-dependent gene expression and metamorphosis in Xenopus laevis. Environmental Health Perspectives 110, 1199e1205. Datta, S., Hansen, L., McConnell, L., Baker, J., LeNoir, J., Seiber, J.N., 1998. Pesticides and PCB contaminants in fish and larvae from the Kaweah River Basin, California. Bulletin of Environmental Contamination and Toxicology 60, 829e836. Davies, P.E., 1987. Physiological, anatomic and behavioural changes in the respiratory system of Salmo gairdneri rich. on acute and chronic exposure to chlorothalonil. Comparative Biochemistry and Physiology, Part C: Comparative Pharmacology 88, 113e119. Davies, P.E., White, R.W.G., 1985. The toxicology and metabolism of chlorothalonil in fish. I. Lethal levels for Salmo gairdneri, Galaxias maculates, G. truttaceus, and G. auratus and the fate of 14C-TCIN in S. gairdneri. Aquatic Toxicology 7, 93e105. Edginton, A.N., Rouleau, C., Stephenson, G.R., Boermans, H.J., 2007. 2,4-D butoxyethyl ester kinetics in embryos of Xenopus laevis: the role of the embryonic jelly coat in reducing chemical absorption. Archives of Environmental Contamination and Toxicology 52, 113e120. Ernst, W., Doe, K., Jonah, P., Young, J., Julien, G., Hennigar, P., 1991. The toxicity of chlorothalonil to aquatic fauna and the impact of its operational use on a pond ecosystem. Archives of Environmental Contamination and Toxicology 21, 1e9. Hashimoto, Y., Nishiuchi, Y., 1981. Establishment of bioassay methods for the evaluation of acute toxicity of pesticides to aquatic organisms. Journal of Pesticide Science 6, 257e264. Koss, R., Wakeford, B., 2000. Rearing Xenopus laevis life history stages. In: Karcher, S.J. (Ed.), Tested Studies for Laboratory Teaching, Proceedings of the 22nd Workshop/Conference of the Association for Biology Laboratory Education (ABLE), vol. 22, pp. 387e393. McMahon, T.A., Halstead, N.T., Johnson, S., Raffel, T.R., Romansic, J.M., Crumrine, P.W., Boughton, R.K., Martin, L.B., Rohr, J.R., 2011. The fungicide chlorothalonil is nonlinearly associated with corticosterone levels, immunity, and mortality in amphibians. Environmental Health Perspectives 119, 1098e 1103.
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Menon, J., Gardner, E.E., Vail, S., 2000. Developmental implications of differential effects of calcium in tail and body skin of anuran tadpoles. Journal of Morphology 244, 31e43. Montforts, M.H.M.M., 1999. Chlorothalonil. CSR Advisory Report 06105A00. Nakajima, K., Fujimoto, K., Yaoita, Y., 2005. Programmed cell death during amphibian metamorphosis. Seminars in Cell & Developmental Biology 16, 271e280. Nieuwkoop, P.D., Faber, J., 1975. Normal Table of Xenopus laevis Development (Daudin): a Systematical and Chronological Survey of the Development from the Fertilized Egg until the End of Metamorphosis, second ed. North-Holland Publishing Company, Amsterdam. Potter, T.L., Wauchope, R.D., Culbreath, A.K., 2001. Accumulation and decay of chlorothalonil and selected metabolites in surface soil following foliar application to peanuts. Environmental Science and Technology 35, 2634e2639.
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Environmental Pollution journal homepage: www.elsevier.com/locate/envpol
Short communication
Effects of chlorothalonil on development and growth of amphibian embryos and larvae Shuangying Yu a, Mike R. Wages a, George P. Cobb b, Jonathan D. Maul a, * a b
Department of Environmental Toxicology, The Institute of Environmental and Human Health, Texas Tech University, Lubbock, TX 79416, USA Department of Environmental Science, Baylor University, Waco, TX 76798, USA
a r t i c l e i n f o
a b s t r a c t
Article history: Received 3 January 2013 Received in revised form 15 May 2013 Accepted 15 June 2013
Chlorothalonil is a broad spectrum fungicide widely used in agricultural and urban environments, yet little is known regarding its effects on amphibians. We examined effects of chlorothalonil on growth, malformations, and mortality in embryos and larvae of Xenopus laevis and Spea multiplicata, and assessed variation in sensitivity among aquatic organisms using a species sensitivity distribution (SSD). Chlorothalonil induced gut malformations in X. laevis embryos and inhibited growth. Tail degeneration was observed in larvae of both species and reduced tail length to total length ratios occurred at environmentally relevant concentrations (5.9 and 11.0 mg/L). The mechanism of tail degeneration is unclear, but alteration in the expression of genes involved in tail resorption is a hypothesized mechanism. Larval amphibians were more sensitive than invertebrates and fish. Based on our results and the range of reported environmental concentrations, chlorothalonil may pose a risk to larval amphibians in certain habitats and scenarios. Ó 2013 Elsevier Ltd. All rights reserved.
Keywords: African clawed frog Fungicide Malformation Mexican spadefoot toad Tail degeneration
1. Introduction Chlorothalonil (2,4,5,6-tetrachloroisophthalonitrile) is the most commonly used fungicide in the U.S. (USEPA, 2011) and is widely used in agriculture to protect crops (USEPA, 1999). Chlorothalonil may be introduced to aquatic habitats by direct application, spray drift, and runoff, which may pose a risk to aquatic organisms. Chlorothalonil concentrations in runoff ranged from 50 to130 mg/L after two days of rainfall (Potter et al., 2001) and can be as high as 500 mg/L during the first runoff event (Wilson et al., 2010). Average body residue in Pacific treefrog (Hyla regilla) tadpoles collected from the Kaweah River basin, CA ranged from 33.3 to 47.7 ng/g wet weight (Datta et al., 1998). Chlorothalonil is highly toxic to fish and invertebrates (USEPA, 1999). The 24e96 h LC50s ranged from 12 to 195 mg/L for freshwater invertebrates and 16 to 76 mg/L for freshwater fish (van Wezel and van Vlaardingen, 2004). Chlorothalonil is toxic to tunicate blood cells at 1 mM (Cima et al., 2008) and fish phagocytes at 250 mg/L (Baier-Anderson and Anderson, 1998). Chronic exposure at 2.0 mg/L can decrease hematocrit and cause gill damage in fish (Davies, 1987). However, studies that address chlorothalonil toxicity to amphibians are limited. A 48-h LC50 of 160 mg/L was reported for * Corresponding author. E-mail address: [email protected] (J.D. Maul). 0269-7491/$ e see front matter Ó 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.envpol.2013.06.017
Japanese common toad Bufo bufo japonicus tadpoles (Hashimoto and Nishiuchi, 1981). Recently, adverse effects on survival, liver tissue and immune cells, and corticosterone concentrations in anurans have been reported at chlorothalonil concentrations between 0.0164 and 164 mg/L (McMahon et al., 2011). Taken together, these studies indicate that chlorothalonil may have a significant impact on amphibians and toxicity data are needed. The objectives of this study were to: (1) examine effects of chlorothalonil on development and growth in two anuran species, African clawed frogs (Xenopus laevis) and Mexican spadefoot toads (Spea multiplicata), and (2) compare sensitivity of larval amphibians and other aquatic organisms to chlorothalonil. 2. Materials and methods 2.1. Experiment 1: embryo and larval Xenopus laevis 96-h acute toxicity tests Breeding, housing, and field collection methods for X. laevis and S. multiplicata have been described in the supplemental materials. Toxicity tests were conducted following standardized FETAX methods (ASTM, 2004) with slight modifications. Embryos were not dejellied to achieve environmental realism and because egg jelly may protect embryos from ambient contaminants (Edginton et al., 2007). Embryos were collected from three clutches, mixed, and examined for viability and developmental stage. Twenty NF stage 8e11 embryos (Nieuwkoop and Faber, 1975) were randomly assigned to each of 50 ml glass jars containing 40 ml FETAX solution spiked with chlorothalonil. Concentrations used were solvent control (acetone), 7.8, 13.0, 21.6, 36.0, and 60.0 mg/L (see Supplemental materials for rationale for choosing these concentrations). There were five replicates for each treatment.
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In the larval experiment, NF stage 46 larvae from three clutches were mixed and 10 larvae were randomly placed in 1 L glass jars containing 900 ml FETAX solution. Treatments included solvent control, 10.2, 12.8, 16.0, 20.0, and 25.0 mg/L, and there were five replicates for each treatment. To compare growth and other morphological measurement before and after exposure, 18 larvae were randomly collected from the same clutches prior to the exposure. These were used to quantify initial size of larvae. Larvae were fed approximately an hour before water change with a diet described in Koss and Wakeford (2000). Experiment 1 was conducted under static renewal conditions with daily water change. All animals were maintained on a 12:12, light:dark cycle at temperatures ranging from 22 to 24 C. Water quality was measured every other day. Embryos and larvae were checked daily for mortality (i.e., no response to prodding). At the end of the exposure, larvae were euthanized and preserved in formalin. Each individual was examined for abnormalities under a microscope. Images of each larvae were taken and total length (head to tail) (TL), body length (BL), and snout-to-vent length (SVL) were measured using Image J (version 1.44P, Wayne Rasband, National Institutes of Health, USA). Larvae exposed to chlorothalonil showed reduced tail length; therefore, tail length was also examined and calculated from TL and BL (Tail ¼ TL BL). We used the tail to TL ratio to minimize the effect of growth rate on absolute tail length measurements and compared the ratio among treatments. Initial larval size measurements prior to the exposure were included in comparisons to demonstrate growth in controls during the experimental period. 2.2. Experiment 2: verification experiments for Xenopus laevis To verify the results of Experiment 1, partial toxicity tests with three replicates per treatment were conducted for embryos and larvae with the same procedures and conditions as previous experiments. However, embryos from only two clutches were used due to lack of breeding or low quality eggs in other clutches. Three clutches were used for the larval experiment. Thirty-one larvae were euthanized and preserved in formalin before the exposure and used to determine initial size measurements. Larvae were fed daily 2 mg of SeraÒ Micro per experimental unit. 2.3. Experiment 3: larval Spea multiplicata 96-h acute toxicity test Larvae (Gosner stage 25) from five clutches were mixed and were randomly assigned to each treatment. Five larvae were placed in 500 ml glass jars with 400 ml laboratory water and six replicates were used for each treatment. A 96-h static non-
renewal toxicity test was performed. Treatments included solvent control, 3.2, 5.4, 9.0, 15.0, and 25.0 mg/L. Fifteen larvae were collected for initial size measurements. Larvae were not fed during the exposure. Water quality was measured at the end of the exposure. Morphological measurements were similar to X. laevis. 2.4. Comparison of chlorothalonil toxicity to amphibians, fish, and invertebrates A species sensitivity distribution (SSD) curve was created for fish, invertebrates, and amphibians to compare sensitivity among taxa and evaluate the risk chlorothalonil poses to aquatic organisms. The SSD curve was fit with the log-probit distribution using the U.S. EPA Species Sensitivity Distribution Generator (SSD_Generator_V1.xlt). We only used 96-h LC50s to minimize the variation due to exposure duration. 2.5. Chemical and statistical analyses Chlorothalonil exposure concentrations were quantified using a gas chromatograph equipped with an electron capture detector. Measured chlorothalonil concentrations were reported throughout with correction for extraction efficiency. Data were analyzed using JMP Statistical Analysis Software (Ver 9.0.0, Statistical Analysis System Institute, Cary, NC, USA) and results were considered significant at a ¼ 0.05 (see Supplemental materials for more information on chemical and statistical analyses).
3. Results and discussion 3.1. Chemical analyses Measured water concentrations of chlorothalonil ranged from 58 to 142% of the nominal concentrations with an average of 96% (Table 1). In Experiment 3, the mean chlorothalonil concentrations were 84, 84, and 76% of the initial concentrations for 3.2, 9.0, and 15.0 mg/L (nominal) treatments, respectively, after 96 h (data provided in Supplemental materials). Water quality data were also provided in Supplemental Materials.
Table 1 Nominal and measured chlorothalonil concentrations (mg/L) at the beginning of exposures, percent mortality at each concentration, 96-h median lethal concentrations (LC50s, 95% confidence interval), and model parameters associated with LC50 estimates for Xenopus laevis and Spea multiplicata. Species
Stage
Nominal (mg/L)
Measured (mg/L)
Mortality (%)
LC50 (95% CI)
Slope
Intercept
X. laevisa
Embryo
Control 7.8 13.0 21.6 36.0 60.0 Control 10.2 12.8 16.0 20.0 25.0
0 8.8 12.8 19.3 36.4 60.7 0 5.9 8.3 14.4 17.0 23.8
0 8 3 6 16 99 10 29 52 82 98 100
42.4 (39.8e45.3)
0.13
5.36
8.2 (7.1e9.2)
0.31
2.57
Control 7.8 13.0 21.6 36.0 60.0 Control 10.2 12.8 16.0 20.0 25.0
0 6.1 9.5 18.8 26.8 60.3 0 8.6 11.1 13.4 16.3 22.5
0 5 2 23 75 100 3 10 0 27 86 100
22.9 (21.4e24.6)
0.25
5.68
14.4 (13.6e15.4)
0.56
8.02
Control 3.2 5.4 9.0 15.0 25.0
0 4.6 7.0 11.0 20.5 34.6
0 0 0 7 93 100
10.7 (9.7e12.2)
0.58
6.26
Larvae
X. laevisb
Embryo
Larvae
S. multiplicata
a b
Larvae
Experiment 1. Experiment 2, verification experiment.
S. Yu et al. / Environmental Pollution 181 (2013) 329e334
3.2. Effects of chlorothalonil on survival The LC50s (95% CI) for X. laevis embryos in Experiment 1 and Experiment 2 were 42.4 mg/L (39.8e45.3) and 22.9 mg/L (21.4e24.6), respectively (Table 1). For X. laevis larvae, the LC50 was 8.2 mg/L (7.1e9.2) in Experiment 1 and 14.4 mg/L (13.6e15.4) in Experiment 2. The LC50 for S. multiplicata larvae was 10.7 mg/L (9.7e12.2), similar to those for X. laevis larvae. Chlorothalonil was less toxic to X. laevis embryos compared to X. laevis larvae, indicating differential sensitivity among developmental stages. Our 96-h LC50s for larvae of both species were approximately 10e20 times lower than the 48-h LC50 (160 mg/L) reported by Hashimoto and Nishiuchi (1981) for B. bufo japonicus, suggesting large variation in sensitivity to chlorothalonil among amphibian species. It should be noted, however, that exposure time varied between our study and that of Hashimoto and Nishiuchi (1981). 3.3. Effects of chlorothalonil on development Chlorothalonil produced malformations in X. laevis embryos, including gut malformations, axial/tail malformations, and edema. Proportions of malformed embryos were significantly different among treatments in both Experiment 1 (ANOVA, F4,20 ¼ 15.5, P < 0.001) and Experiment 2 (F4,10 ¼ 24.7, P < 0.001) (Fig. 1).
100
A
%Total malformations %Gut malformatinons %Axial/tail malformations %Edema
%Malformation
80
60
** 40
20
331
Compared to controls (mean SE: 4.92 2.20% and 9.24 2.50% in Experiment 1 and 2, respectively), incidence of total malformations was significantly higher at 36.4 mg/L (42.56 5.02%) in Experiment 1 and at 18.8 mg/L (41.11 8.04%) and 26.8 mg/L (93.33 6.67%) in Experiment 2 (Dunnett’s test, all P < 0.05). Chlorothalonil at 26.8 and 36.4 mg/L produced significantly greater proportions of gut malformations and/or axial/tail malformations compared to controls (all P < 0.05) (Fig. 1). Most interestingly, X. laevis larvae exhibited tail shortening during chlorothalonil exposure (Fig. 2). Because of the high mortality at the two highest concentrations (i.e., 100% mortality or data from only one replicate), these two treatments were not included in statistical analyses. Tail to TL ratios were significantly different among treatments in Experiment 1 (KruskaleWallis test, P < 0.001), and tail to TL ratios for all chlorothalonil concentrations (0.56 0.006) were significantly lower compared to control (0.63 0.002) and initial (0.64 0.003) (all P < 0.001) (Fig. 3A). In Experiment 2, larvae from 11.1 (0.61 0.007) and 13.4 mg/L (0.56 0.006) had significantly lower tail to TL ratios than control (0.66 0.001) and initial (0.67 0.004) (all P < 0.01) (Fig. 3B). Similar to X. laevis, we observed tail shortening in S. multiplicata, and larvae at 11.0 (0.57 0.003) and 20.5 mg/L (0.37 0.063) had significantly lower tail/TL ratio compared to control (0.60 0.002) (all P < 0.05) (Fig. 3C). Tail degeneration caused by chlorothalonil in both species is the most important finding of this study. External Ca2þ has been reported to induce apoptosis and tail shortening in tadpoles (Menon et al., 2000) and acetochlor has been demonstrated to reduce tadpole body area and alter expression of T3 dependent genes (Crump et al., 2002). But to our knowledge, no study has reported that pesticides or other chemicals cause such remarkable tail shortening in early larval stages (i.e., NF stage 46 or Gosner stage 25). Both X. laevis and S. multiplicata larvae exposed to chlorothalonil had shorter tails compared to the initial group from the same clutches collected prior to the exposure. This difference suggests that the observed tail shortening is likely a degeneration process rather than growth inhibition. The mechanism of regressing tails is unknown but may be related to apoptosis of tail tissue similar to tail absorption during metamorphosis (Nakajima et al.,
0 Control
8.8
12.8
19.3
36.4
B
**
100
%Malformation
80
60
* *
40
20
0 Control
6.1
9.5
18.8
26.8
Concentration (µg/L) Fig. 1. Incidence of total, gut, and axial/tail malformations and edema in stage 8e11 Xenopus laevis embryos exposed to chlorothalonil for 96 h in Experiment 1 (A) and Experiment 2 (B). Concentrations on the x-axis are measured concentrations. Error bars are SE and asterisks indicate a significant difference compared to controls (P < 0.05).
Fig. 2. Tail shortening in representative stage 46 Xenopus laevis larvae exposed to different concentrations of chlorothalonil for 96 h in Experiment 1 (from top to bottom: control, 5.9, 8.3, and 14.4 mg/L).
0.70
0.70
A
B
0.65
0.65
0.60
0.60
Tail/TL ratio
Tail/TL ratio
* * *
0.55
*
0.50 0.45
*
0.55 0.50 0.45
0.40
0.40 Initial
Control
5.9
8.3
14.4
17
Initial
Control
8.6
11.1
13.4
16.3
Concentration (µg/L)
Concentration (µg/L)
0.65
C
0.60
*
Tail/TL ratio
0.55 0.50 0.45
*
0.40 0.35 0.30 Initial Control
4.6
7
11
20.5
Concentration (µg/L)
Fig. 3. Tail/total length (TL) ratio of stage 46 Xenopus laevis larvae exposed to chlorothalonil for 96 h in Experiment 1 (A) and Experiment 2 (B); and Gosner stage 25 Spea multiplicata larvae exposed to chlorothalonil for 96 h in Experiment 3 (C). The initial group represents morphological measurements on a haphazardly selected subset of tadpoles preserved immediately prior to the exposure. There was only one replicate remaining at 17.0 mg/L in Experiment 1 and 16.3 mg/L in Experiment 2. Concentrations on the x-axis are measured concentrations. Error bars are SE and asterisks indicate a significantly different tail/TL ratio compared to controls (P < 0.05). 4.0
4.0
B
3.8
3.8
3.6
3.6
SVL (mm)
SVL (mm)
A
* 3.4
*
3.4
3.2
3.2
3.0
3.0
*
Control
8.8
12.8
19.3
36.4
Control
Concentration (µg/L)
C
* *
*
18.8
26.8
3.5
D
5.0
SVL (mm)
SVL (mm)
*
4.0
9.5
Concentration (µg/L)
5.0
4.5
6.1
4.5
* *
4.0
3.5
Initial
Control
5.9
8.3
14.4
Concentration (µg/L)
17
Initial
Control
8.6
11.1
13.4
16.3
Concentration (µg/L)
Fig. 4. Snout-to-vent length (SVL) of stage 8e11 Xenopus laevis embryos in Experiment 1 (A) and Experiment 2 (B) and stage 46 Xenopus laevis larvae in Experiment 1 (C) and Experiment 2 (D). Both experiments were 96 h chlorothalonil exposures. Concentrations on the x-axis are measured concentrations. Error bars are SE and asterisks indicate a significantly different SVL compared to controls (P < 0.05).
S. Yu et al. / Environmental Pollution 181 (2013) 329e334
2005). It is possible that chlorothalonil may interfere with pathways that trigger tail resorption by altering expression of genes involved in tail cell death (Brown et al., 1996) (see Supplemental materials for additional discussion).
333
greatest sensitivity to chlorothalonil and that additional amphibian species may be needed to fully evaluate the risk this fungicide may pose to amphibian populations. 4. Conclusions
3.4. Effects of chlorothalonil on growth There was a significant difference in SVL among treatments for X. laevis embryos in both Experiment 1 (F4,20 ¼ 10.4, P < 0.001) (Fig. 4A) and Experiment 2 (F4,10 ¼ 29.9, P < 0.001) (Fig. 4B). Embryos from 36.4 mg/L (3.42 0.04 mm) in Experiment 1 and embryos from 18.8 mg/L (3.53 0.09 mm) and 26.8 mg/L (3.03 0.04 mm) in Experiment 2 were smaller than controls (3.64 0.04 and 3.77 0.04 mm for Experiment 1 and 2, respectively, all P < 0.001). Xenopus laevis larvae collected from the initial group and all chlorothalonil concentrations (the two highest concentrations not included) were significantly smaller (4.43 0.05 mm) than larvae from the control treatment (4.94 0.06 mm, all P < 0.001) in Experiment 1 (Fig. 4C). In Experiment 2, larvae from the initial group (4.22 0.09 mm) and 13.4 mg/L (3.87 0.08 mm) were significantly smaller compared to the control group (4.96 0.01 mm, all P < 0.001) (Fig. 4D). For S. multiplicata larvae, only larvae from the initial group had smaller SVL (3.26 0.09 mm) compared to the control (4.16 0.04 mm, P < 0.001) (see Supplemental materials for the figure and additional discussion). 3.5. Comparison of chlorothalonil toxicity to amphibians, fish, and invertebrates The 96-h LC50s for fish and invertebrates were obtained from the U.S. EPA OPP Pesticide Ecotoxicity Database and existing literature (Davies and White, 1985; Ernst et al., 1991; Montforts, 1999; Sherrard et al., 2002) (data provided in Supplemental materials). Spea multiplicata and X. laevis were more sensitive than fish and invertebrates that had available 96-h LC50 data (Fig. 5). The estimated 5% hazard concentration (HC5, 95% CI) was 6.28 mg/L (3.52e 11.20), and the HC95 was 178.55 mg/L (100.06e318.59). When the amphibian LC50s were compared among taxa of freshwater organisms, the LC50s of X. laevis and S. multiplicata larvae were near the lower range of LC50s for fish, indicating a greater sensitivity to chlorothalonil. Comparisons among all freshwater taxa suggests that larval amphibians may have the
Proportion of species affected
1.2 Predicted relationship 95% Confidence interval Amphibians Fish Invertebrates
1.0
0.8
Ceriodaphnia dubia Ictalurus punctatus Lepomis macrochirus Cyprinus carpio Oncorhynchus mykiss
0.6
Galaxia auratus Pimephales promelas
0.4
Galaxia truttaceus Galaxia maculatus Paratya australiensis Astacopsis gouldi Xenopus laevis Spea multiplicata
0.2
0.0
1
10
100
1000
96-h LC50 (µg/L) Fig. 5. Species sensitivity distribution (SSD) for chlorothalonil among freshwater fish, invertebrates, and data from two amphibian species, Xenopus laevis and Spea multiplicata from the present study. All data used in the SSD are 96-h median lethal concentrations (i.e., LC50s).
Our results suggest that chlorothalonil is toxic to X. laevis and S. multiplicata, and X. laevis larvae were more sensitive to chlorothalonil than embryos. Our study reveals for the first time that chlorothalonil disrupts tail development and causes tail degeneration. Other sublethal effects include inhibition of growth and malformations. Based on acute toxicity data, amphibians appear to be more sensitive to chlorothalonil and may be at greater risk compared to other aquatic organisms. Further investigation focused on molecular and cellular pathways are needed to reveal the mechanism of tail shortening observed in the present study. Acknowledgments S Yu received funding from the Paul Whitfield Horn Fellowship provided by the University Women’s Club of Texas Tech University. Appendix A. Supplementary data Supplementary data related to this article can be found at http:// dx.doi.org/10.1016/j.envpol.2013.06.017. References American Society for Testing and Materials, 2004. Standard Guide for Conducting the Frog Embryo Teratogenesis Assay-Xenopus (FETAX). ASTM E1439e98 (2004). Baier-Anderson, C., Anderson, R.S., 1998. Evaluation of the immunotoxicity of chlorothalonil to striped bass phagocytes following in vitro exposure. Environmental Toxicology and Chemistry 17, 1546e1551. Brown, D.D., Wang, Z., Furlow, J.D., Kanamori, A., Schwartzman, R.A., Remo, B.F., Pinder, A., 1996. The thyroid hormone-induced tail resorption program during Xenopus laevis metamorphosis. Proceedings of the National Academy of Sciences 93, 1924e1929. Cima, F., Bragadin, M., Ballarin, L., 2008. Toxic effects of new antifouling compounds on tunicate haemocytes I. Sea-Nine 211 and chlorothalonil. Aquatic Toxicology 86, 299e312. Crump, D., Werry, K., Veldhoen, N., Van Aggelen, G., Helbing, C.C., 2002. Exposure to the herbicide acetochlor alters thyroid hormone-dependent gene expression and metamorphosis in Xenopus laevis. Environmental Health Perspectives 110, 1199e1205. Datta, S., Hansen, L., McConnell, L., Baker, J., LeNoir, J., Seiber, J.N., 1998. Pesticides and PCB contaminants in fish and larvae from the Kaweah River Basin, California. Bulletin of Environmental Contamination and Toxicology 60, 829e836. Davies, P.E., 1987. Physiological, anatomic and behavioural changes in the respiratory system of Salmo gairdneri rich. on acute and chronic exposure to chlorothalonil. Comparative Biochemistry and Physiology, Part C: Comparative Pharmacology 88, 113e119. Davies, P.E., White, R.W.G., 1985. The toxicology and metabolism of chlorothalonil in fish. I. Lethal levels for Salmo gairdneri, Galaxias maculates, G. truttaceus, and G. auratus and the fate of 14C-TCIN in S. gairdneri. Aquatic Toxicology 7, 93e105. Edginton, A.N., Rouleau, C., Stephenson, G.R., Boermans, H.J., 2007. 2,4-D butoxyethyl ester kinetics in embryos of Xenopus laevis: the role of the embryonic jelly coat in reducing chemical absorption. Archives of Environmental Contamination and Toxicology 52, 113e120. Ernst, W., Doe, K., Jonah, P., Young, J., Julien, G., Hennigar, P., 1991. The toxicity of chlorothalonil to aquatic fauna and the impact of its operational use on a pond ecosystem. Archives of Environmental Contamination and Toxicology 21, 1e9. Hashimoto, Y., Nishiuchi, Y., 1981. Establishment of bioassay methods for the evaluation of acute toxicity of pesticides to aquatic organisms. Journal of Pesticide Science 6, 257e264. Koss, R., Wakeford, B., 2000. Rearing Xenopus laevis life history stages. In: Karcher, S.J. (Ed.), Tested Studies for Laboratory Teaching, Proceedings of the 22nd Workshop/Conference of the Association for Biology Laboratory Education (ABLE), vol. 22, pp. 387e393. McMahon, T.A., Halstead, N.T., Johnson, S., Raffel, T.R., Romansic, J.M., Crumrine, P.W., Boughton, R.K., Martin, L.B., Rohr, J.R., 2011. The fungicide chlorothalonil is nonlinearly associated with corticosterone levels, immunity, and mortality in amphibians. Environmental Health Perspectives 119, 1098e 1103.
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Sherrard, R.M., Murray-Gulde, C.L., Rodgers Jr., J.H., Shah, Y.T., 2002. Comparative toxicity of chlorothalonil and chlorpyrifos: Ceriodaphnia dubia and Pimephales promelas. Environmental Toxicology 17, 503e512. United States Environmental Protection Agency, 1999. Re-registration Eligibility Decision. Chlorothalonil, Washington, DC. United States Environmental Protection Agency, 2011. Pesticide Industry Sales and Usage: 2006 and 2007 Market Estimates. Available online: http://www.epa.gov/ opp00001/pestsales/07pestsales/market_estimates2007.pdf. van Wezel, A.P., van Vlaardingen, P., 2004. Environmental risk limits for antifouling substances. Aquatic Toxicology 66, 427e444. Wilson, P.C., Riiska, C., Albano, J.P., 2010. Nontarget deposition and losses of chlorothalonil in irrigation runoff water from a commercial foliage plant nursery. Journal of Environmental Quality 39, 2130e2137.