In vivo genotoxicity of atrazine to anuran larvae

Mutation Research 560 (2004) 69–78

In vivo genotoxicity of atrazine to anuran larvae J.L. Freeman, A.L. Rayburn∗ Department of Crop Sciences, 320 ERML, 1201 West Gregory, University of Illinois, Urbana, IL 61801, USA Received 17 September 2003; received in revised form 19 February 2004; accepted 19 February 2004

Abstract Atrazine has been an environmental contaminant for more than two decades. While there can be little dispute as to the presence of atrazine in non-target watersheds, the debate has centered on the consequences of this contamination. The purpose of this study was to determine if atrazine is genotoxic to developing anurans. Anurans are one of the groups that have the highest potential for being affected by watershed contamination. In initial studies, larvae from two anuran species were exposed to known genotoxic agents. Upon flow cytometric analysis, those organisms exposed to the genotoxic agents resulted in a statistically significant increase in nuclear heterogeneity. Having demonstrated that flow cytometric analysis could be used to detect genotoxicity in anuran larvae, the larvae of the two species were exposed to different levels of atrazine for various durations. The concentrations and lengths of exposure were consistent (albeit on the higher side) with conditions found in the Midwestern US. In neither species was an increase in nuclear heterogeneity observed. Thus, atrazine at levels and time of exposure representing conditions found contaminating Midwestern watersheds does not appear to be genotoxic to developing anurans. © 2004 Published by Elsevier B.V. Keywords: Toads; Xenopus; Genotoxicity; Flow cytometry; Atrazine

1. Introduction Contamination of watersheds by pesticides is an important issue in United States agriculture. Taylor [1] reported that herbicide contamination could be found in many waterways in Illinois. Constant exposure to toxic chemicals over the life span of organisms has been speculated to have cumulative deleterious effects to these organisms. Amphibian species such as anurans are especially vulnerable due to ova being fertilized in the water where the embryos develop into tadpoles and later metamorphose into adults. Embryogenesis and organogenesis take place in the aquatic environ∗ Corresponding author. Tel.: +1-217-333-4374; fax: +1-217-333-4582. E-mail address: [email protected] (A.L. Rayburn).

1383-5718/$ – see front matter © 2004 Published by Elsevier B.V. doi:10.1016/j.mrgentox.2004.02.008

ment amidst the herbicide contamination. Presence of herbicides in the water may have adverse effects on amphibian populations. Amphibian populations have been reported to be declining at rates exceeding other vertebrates. Unlike many other environmental observations and concerns such as global warming, the decline of amphibian populations is universally accepted. Suspected causes include habitat destruction, UV exposure, infectious diseases, and pollutants, including those that alter sexual development [2]. Amphibians, which have lives dependent upon water and land, fulfill essential roles in the environment as both predator and prey, and thus are unique sentinels of ecosystem health. As herbicide contamination increases in wetland ecosystems, a deterioration of the ecosystem may occur. Levels of the herbicide atrazine have been reported to be up to

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860 ppb in streams below field plots that have been treated with atrazine [3]. Tailwater pits in Kansas have been reported to have a mean atrazine contamination level of 50–100 ppb with concentrations as high as 1 ppm [4]. In addition to atrazine, levels of various other agrochemicals were also reported. Thus, in the Midwest, agrochemical contamination of aquatic ecosystems is not uncommon and is in fact the norm. The most well characterized and widely used agrochemical that is known to contaminate surface waters is atrazine. The mode of action of atrazine is blocking electron transport in photosystem II leading to chlorophyll destruction and blocking photosynthesis [5]. When atrazine was first released for agricultural use it was thought that since photosynthesis is limited to plants, animals would be immune to any effects of atrazine. It was soon suspected that atrazine might have non-target action in animals. Atrazine has been implicated as a clastogen (an agent that causes chromosomal damage). Yoder et al. [6] found an increase of chromosomal aberrations in lymphocyte cultures of farm workers exposed to atrazine. Atrazine was shown to cause whole cell clastogenicity and to induce chromosome breakages in CHO cells through flow cytometric analysis at low concentrations [7–10]. Ribas et al. [11] also demonstrated that atrazine could be genotoxic to animal chromosomes. Atrazine has been reported to have significant affects on frogs with increased mortality seen in leopard frog (Rana pipens) tadpoles exposed to atrazine for 27 days at concentrations as low as 310 ppb [12]. Clements et al. [13] observed that bullfrog tadpoles exposed to atrazine for 24 h had significant DNA damage. Reeder et al. [14] found an association between the detection of atrazine and intersex cricket frogs (Acris crepitans). Diana et al. [15] reported that atrazine exposure could lead to reduction in fitness of wild populations of anurans. Recently, Hayes et al. [16] reported that exposure levels as low as 0.1 ppb resulted in hermaphroditism in Xenopus laevis larvae under laboratory conditions. Thus, exposure to atrazine has been reported to be deleterious to amphibian species at several endpoints that could affect the survival of a species. If a mutagen has varying effects on the DNA content of a population, the distribution of nuclei with respect to DNA content will appear quite broad due to

increasing nuclear heterogeneity. Thus, the coefficient of variation (CV) of the distribution will be large. Several investigators have reported the phenomenon of increasing CV in response to mutagens. Combinations of increased CVs, cell cycle disruption and aneuploid peaks, as measured by flow cytometry, referred to as whole cell clastogenicity, are often seen [7,10,17–21]. The advantage of whole cell clastogenicity is that it is a quick, reliable technique to efficiently assess the potential genomic damage to cells [9]. Parameters identified that may be associated with the widening of the CV include chromosome structure changes, modifications in chromatin composition, apoptosis and cell cycle changes. While each of these mechanisms is different they do indicate alterations that could have negative consequences. The specific objective of this research was to determine if the levels of atrazine contamination known to occur in watersheds could induce instability in anuran chromosomes. Atrazine was chosen for this research for two reasons. One, it is the most common agrochemical contaminate of watersheds and two, it will serve as a model agrochemical to develop the necessary methodology due to the tremendous amount of information already available about this herbicide. Since X. laevis have been researched extensively, they served as a reference model in this study. The American toad (Bufo americanus) was used to investigate the effects of atrazine on a native amphibian species.

2. Materials and methods 2.1. Positive controls 2.1.1. Bufo americanus Adult B. americanus were collected from the wild in the fall, over-wintered in the laboratory and bred in the spring for experimental use. Tadpoles were housed in 10 gallon tanks under controlled conditions. For chemical exposure tadpoles were placed in 3 l tanks. Ara-C (cytosine ␤-d-arabinofuranoside, CAS No. 147-94-4) was purchased from Sigma (St. Louis, MO). Tadpoles were exposed to 0, 0.3 or 3 mM Ara-C for 72 h under a chemical hood with aeration. The nuclear DNA content distribution was then analyzed. Tadpoles were prepped for nuclear DNA content using a modification of the Gold et al. [22] fish cell

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protocol for flow cytometry. Tadpoles were homogenized in 10 ml of a 1× phosphate buffered saline (PBS) (Sigma) and then filtered through a 53 ␮m mesh. Four 1 ml aliquots of nuclei suspension per sample was placed in microfuge tubes and centrifuged for 8 s. The supernatant were aspirated and nuclei suspensions combined for each sample into one resuspension of 800 ␮l PBS. Centrifugation was repeated, supernatant aspirated and sample resuspended in 800 ␮l of PBS. Nuclei samples were fixed in 50% ice cold EtOH for 20 min at −20 ◦ C. Samples were diluted 1:2000 in isoton solution for nuclei counting. Nuclei counts were performed on a Coulter Counter Z Series (Coulter Electronics, Hileah, FL). Two nuclei counts were taken and averaged for each sample. After counting and fixing, nuclei suspensions were centrifuged for 8 s, washed with PBS and resuspended in the calculated propidium iodide (PI) stain solution [0.8 g sodium chloride (137 mM), 0.056 g PIPES, 0.028 g disodium EDTA (0.75 mM), 4 mg deoxyribonuclease-free ribonuclease and 5 mg propidium iodide in 100 ml distilled water brought to pH 7.50 with NaOH] volume to achieve 6 × 105 nuclei per ml of stain. All samples were placed on ice in the dark for 1 h and filtered through a 53 ␮m mesh. Flow cytometric analysis was done on an EPICS XL flow cytometer (Coulter Electronics, Hileah, FL) using an excitation wavelength of 488 nm. Ten thousand nuclei were analyzed per sample. The coefficient of variation of the G1 peaks were measured and compared. An analysis of variance (ANOVA) was run using SAS. Means were compared using the least significance difference (LSD) test at α = 0.05 when a significant ANOVA was observed. 2.1.2. Xenopus laevis X. laevis tadpoles were obtained from NASCO (Fort Atkinson, WI) and Xenopus Express (Plant City, FL). Tadpoles were housed in 10 gallon tanks under controlled conditions once received in the laboratory. Tadpoles were exposed to 0 or 3 mM Ara-C for 72 h under a chemical hood with aeration in 3 l tanks. After exposure to the treatments, the DNA distribution was analyzed as described in the B. americanus Ara-C exposure. In addition, ethyl methanesulfonate (EMS), CAS No. 62-50-0) was also tested. Tadpoles were exposed to 0, 150 and 300 ppm EMS for 72 h as previously

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described. Two tadpoles were placed in each tank. Every 24 h, the tadpoles were transferred into a new tank with the appropriate treatment. The experimental unit was the tank with two subsamples per tank. The experiment was replicated five times for a total of five tanks per concentration. The nuclei were isolated and analyzed as described above. 2.2. Atrazine 2.2.1. B. americanus Adult B. americanus were collected from the wild in Illinois in the fall, over-wintered in the laboratory and bred in the spring for experimental use. Atrazine (CAS No. 1912-24-9) was purchased from Chem Services, Inc. Tadpoles were housed in 10 gallon tanks under controlled conditions. In the first set of experiments, tadpoles were exposed to 0, 250, 500, 1000, 5000 or 10,000 ppb atrazine in 3 l tanks. Two tadpoles were placed in each tank. Water was changed in the tanks once per week and water samples collected. Water conditions were monitored throughout the experiment. The atrazine concentration of the water was tested using an ELISA concentration kit (Abraxis, Ivyland, PA). After 3 weeks of exposure, two tadpoles (subsamples) per tank were analyzed. Tanks were considered as the experimental unit. DNA content was analyzed as described in the B. americanus Ara-C exposure. In the second set of experiments, tadpoles were exposed to 0 or 800 ppb atrazine for 3 weeks. Once again two tadpoles were placed in each 3 l tank. After 3 weeks of exposure, two tadpoles were analyzed per tank. Experimental conditions and analysis was done as described previously. 2.2.2. X. laevis X. laevis tadpoles were obtained from NASCO (Fort Atkinson, WI). Tadpoles were housed in 10 gallon tanks under controlled conditions once received in the laboratory. In the first experiment, tadpoles were placed in 3 l tanks and exposed to 0 or 800 ppb atrazine. After 3 weeks exposure, two tadpoles per tank were analyzed as described in the B. americanus Ara-C exposure. Experimental conditions were monitored and water samples collected at each week for analysis as described previously.

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In the second set of experiments, tadpoles were exposed to 0 or 800 ppb atrazine. DNA content was analyzed after 3, 4 and 5 weeks of exposure. Flow analysis could not be done after the first 2 weeks of exposure due to the size of the animals being so small that the number of nuclei isolated was insufficient to allow for the analysis. Experimental conditions and analysis was performed as described previously.

3. Results

difference in CV was noted between the treatments (P = 0.0028) (Fig. 2A and B). The mean of 10 replications of the control tadpoles was 6.2 while the mean of the 3 mM Ara-C treatment group was 7.0. With respect to concentration of EMS, the mean CVs of the nuclei were found to be significantly different (P = 0.0427). Both the 150 and 300 ppm EMS resulted in a significantly higher CV than the 0 ppm treatment at α = 0.05 (Fig. 3). While no statistical significance was observed between the 150 and 300 ppm concentrations, a numerical increase in the CV was noted.

3.1. Positive controls 3.2. Atrazine 3.1.1. B. americanus Nuclei isolated from the tadpoles resulted in a histogram with one sharp distinct peak (Fig. 1A). Upon comparison with chicken red blood cells the genome size of B. americanus was calculated to be ≈9 pg per 2C nucleus. This is in the range of genome sizes that have been previously reported for B. americanus [23–25]. Tadpoles that were exposed to different concentrations of Ara-C were observed to have statistically different CVs at P = 0.0001 (Fig. 1B). The eight animals that were not exposed to Ara-C had a mean CV of approximately 8.4. The eight larvae exposed to 0.3 mM Ara-C had a mean CV of 10.6, which was significantly different (α = 0.05) than both the control animals and the animals exposed to the higher concentration. The eight tadpoles exposed to 3 mM Ara-C had a CV of 12.5, which was significantly different (α = 0.05) from both the control and 0.3 mM concentrations (Fig. 1B). Ara-C exposure for 72 h resulted in a dose response with respect to increasing CV from nuclei isolated from animals exposed to increasing Ara-C concentration. 3.1.2. X. laevis Upon comparison with chicken red blood cells as an internal standard (defined as having 2.5 pg/2C [26]), the genome size observed for the X. laevis was 6.3 pg/2C, within the reported range of X. laevis [27]. Since 3 mM Ara-C was observed to give the largest significant increase in CV, this concentration was selected for the X. laevis experiments. Comparing the full peak CVs of 20 control animals versus the 20 tadpoles exposed to 3 mM Ara-C, a significant

3.2.1. B. americanus Initially, a wide range of atrazine concentrations was run to determine if atrazine was genotoxic at any level. Upon water quality analysis, the atrazine levels were observed to be within the expected range of each treatment. Thus, no degradation of atrazine occurred within the 1 week between tank water changes. A 3-week exposure was chosen since this would be the longest period of time that anuran larvae would be exposed to high levels of atrazine due to the seasonal fluctuations of atrazine contamination. The concentrations up to 1000 ppb relate to high exposure levels that can be observed in the field while 5000 and 10,000 ppb far exceed any expected exposure level. As can be seen in Table 1, no significant increase was observed among the different treatments (P = 0.6047). Thus, the indication is that atrazine was not genotoxic to the tadpoles. However, when just the raw numbers were compared, it did appear that overall the CV of the nuclei was increasing. It should be noted that no acute toxicity was noted at any of the atrazine Table 1 CV of nuclei isolated from toad tadpoles exposed to various levels of atrazine Atrazine concentration (ppb)

N

Average full peak CV

Standard deviation

0 250 500 1000 5000 10000

6 6 6 6 6 6

5.7 5.7 5.7 5.5 6.6 6.5

0.7 0.5 0.3 1.1 1.0 1.6

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Fig. 1. Flow histograms of nuclei isolated from B. americanus. (A) Flow histogram of nuclei isolated from a control tadpole; (B) flow histogram of nuclei isolated from a tadpole exposed to 3 mM Ara-C.

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Fig. 2. Flow histogram of nuclei isolated from X. laevis. (A) Flow histogram of nuclei isolated from a control tadpole; (B) flow histogram of nuclei isolated from a tadpole exposed to 3 mM Ara-C.

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7.5

7.0

CV

6.5

6.0

5.5 0.0 0

150

300

Concentration EMS (ppm) Fig. 3. Mean CVs of the X. laevis tadpoles exposed to 0, 150 or 300 ppm EMS increase with dose. The mean CV of the 0 treatment was 6.0, the mean CV of the 150 ppm treatment was 6.7 and the mean CV of the 300 ppm treatment was 6.8. Both EMS treatments were significantly different from the control (*α = 0.05).

concentrations tested. In order to ensure that atrazine was indeed not genotoxic, a second experiment was run. In this experiment, two treatments were selected, 0 and 800 ppb atrazine. The average CV of nuclei isolated from the 0 ppb atrazine treated tadpoles was 5.4 ± 0.3 while the average CV of the nuclei of the tadpoles treated with 800 ppb atrazine was 5.4 ± 0.5. The total number of animals in each treatment was 36. No significant difference was observed (P = 0.1978). 3.2.2. X. laevis In the first experiment, 18 animals were exposed to no atrazine and 18 were exposed to 800 ppb atrazine. The mean CV of the nuclei isolated from the control tadpoles was 5.8 ± 0.7 while the mean CV of the nuclei isolated from the larvae exposed to 800 ppb was 6.3 ± 1.4. The means were not significantly different with a P-value of 0.1756. Although the difference was not significant, the CV of the 800 ppb atrazine was numerically larger. In order to ensure that atrazine exposure did not increase the CV, a second experiment was conducted.

In this second experiment, length of exposure was also varied at the two concentrations tested. After a 3 week exposure, no significant difference in CV was observed between nuclei isolated from the 0 ppb treated tadpoles and the nuclei from 800 ppb treated tadpoles (Table 2, P = 0.1650). In addition, in this experiment the mean CV of the nuclei isolated from the 800 ppb treated tadpoles was numerically lower than the nuclei isolated from the 0 ppb treated tadpoles. After both 4 and 5 week exposures, no significant difference was noted with respect to CV (P = 0.5325 and 0.6952, respectively). Table 2 CVs of the nuclei isolated from X. laevis exposed to 0 or 800 ppb atrazine for various times Concentration of atrazine (ppb)

Weeks exposure

N

Mean CV

0 800

3 3

6 6

6.2 ± 0.4 5.8 ± 0.4

0 800

4 4

6 6

5.0 ± 0.3 5.7 ± 1.2

0 800

5 5

6 6

5.4 ± 0.6 5.6 ± 0.6

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4. Discussion 4.1. Positive controls In both species, Ara-C was observed to result in a significant increase in CV indicating whole cell clastogenecity. Ara-C is a widely used chemotherapy drug in the treatment of acute myeloid leukemia [28,29]. Ara-C is known to interfere with DNA synthesis [30] and is clastogenic [7,31,32]. Ara-C exposure has been reported to result in an increase in the CV of the G1 peak of CHO cells [9,10]. Biradar and Rayburn [7] were able to determine that this increase in CV was due partly to chromosome damage by the use of flow karyotyping. However, in this case, we were not able to determine if the increase in the CV of the Ara-C treated animals was due to chromosome damage or cell cycle disruption. Peralta et al. [33] observed that Ara-C does affect intercellular organization in Bufo arenarum embryos. Therefore, the Ara-C affects observed in the present study could be due to the affect of Ara-C on cell division or chromosome damage. In order to ensure that the response observed was a genotoxic response, a second positive control was examined in X. laevis. EMS is a known clastogen and has been listed as a potential positive control standard in genotoxic assays [34]. Wagner et al. [35] calibrated flow cytometric analysis with the single cell gel electrophoresis assay and forward mutation assay using EMS and found a close correspondence between the three assays with respect to genotoxic endpoints. Thus, EMS affects seen using our flow cytometric assay can be hypothesized to be due to a genotoxic response. To minimize any confounding effects of toxicity and animal metamorphosis, the exposure time was limited to 72 h. No signs of distress or decrease in feeding were observed during the experiment. Placing the tadpoles in fresh treatment every 24 h was due to the reported reduction in genotoxic response associated with EMS degradation in water [36]. The concentration of 150 ppm EMS used in this study was chosen based on the results of Jaylet et al. [37] who observed a positive response with respect to the micronuclei assay in 4 days with a 124 ppm concentration. Due to the shorter exposure time, the EMS concentration used in this study was increased slightly

to 150 ppm. The concentration was then doubled, again due to the shorter exposure time. The results presented here indicate that the increase in CV of larvae nuclei exposed to EMS gives comparable results with the micronuclei assay of Jaylet et al. [37]. The 150 ppm EMS gave a significant increase in the nuclei of exposed larvae compared to the 0 ppm treatment. This is in agreement with other studies with a plethora of organisms that demonstrate a genotoxic response with EMS [35–38]. Observing a positive response at 150 ppm EMS after such a short exposure time testifies to the sensitivity of this technique. Hummelen et al. [36] hypothesized since EMS is considered to be a weak clastogen that obtaining a positive clastogenic response with this chemical is a “good indication of the sensitivity” of genotoxic assays. Flow cytometry was successful in determining the effects of the genotoxic agents Ara-C and EMS on anuran tadpoles. The next step was to determine if the ubiquitous agrochemical contaminant atrazine is genotoxic to tadpole larvae. 4.2. Atrazine Whether atrazine is an environmental concern has been debated for well over two decades. Since the 1980’s published reports using different techniques have resulted in conflicting data with respect to the potential genotoxicity of atrazine [7,8,10,39–43]. While the debate over the potential genotoxicty of atrazine continues, the major emphasis of the debate seems to be shifting. The question now being addressed is: do any potential genotoxic effects of atrazine have any relevance in risk assessment? Tennant et al. [44] observed, using the alkaline single cell gel assay, that atrazine did indeed induce DNA damage but that the concentration necessary to induce damage was so high that it had no environmental relevance. Rayburn et al. [9] using flow cytometry reached a similar conclusion. Although they did observe DNA damage induced by atrazine at what could be termed environmental levels, upon comparison with the DNA damage induced with caffeine, an approved food substance, relevant levels of exposure of caffeine induced more damage that that of atrazine. The purpose of this study was to determine whether the highest concentration of atrazine at the longest

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possible exposure time would be genotoxic to anuran larvae. The 800 ppb atrazine level was selected because it was less than the highest levels that have been found polluting waterways and more accurately represented a concentration one may routinely find in a highly contaminated area. The higher concentrations of 5000 and 10,000 ppb may be present in a spill area but these concentrations would be much higher than one might expect from runoff of a treated field. The native species, B. americanus, was used to ensure that any affect would be relevant to the natural environment. X. laevis was used due to its well documented use as a model amphibian species. Since no genetic damage was observed in either species it seems most logical that atrazine is not genotoxic to most anuran larvae. This is not to dismiss the possibility of some species being highly sensitive to atrazine. This also does not mean that atrazine could not be causing DNA damage. Such damage could be repaired by the organism and thus would not represent an imminent threat to the species. In addition, atrazine could be an environmental threat at an endpoint other than the endpoint measured in this study. However, with respect to DNA damage as measured in this study, atrazine does not appear to be an environmental threat to amphibians.

Acknowledgements We thank Dr. B. Pilas of the flow cytometry facility of the Biotechnology Center, University of Illinois for her assistance and Dr. Kimberlee Beckmen, Department of Veterinary Biosciences, College of Veterinary Medicine, University of Illinois for providing the American toad tadpoles. We also thank the Interdisciplinary Environmental Toxicology Program, University of Illinois for awarding JLF as an Environmental Toxicology Scholar. This research was partially funded by Grant numbers IL-97-3; IL-99-5 from the Illinois Water Resource Center, Dr. V. Beasley (P.I.), Dr. D. Bunick (Co-P.I.) and Dr. A.L. Rayburn (Co-P.I.). This material is based upon work supported by the USDA-Hatch under Award No. ILLU-15-0389. Any opinions, findings and conclusions or recommendations expressed in this publication are those of the author(s) and do not necessarily reflect the views of the USDA-Hatch.

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