Tyrone Hayes, Kelly Haston, Mable Tsui, Anhthu Hoang, Cathryn Haeffele, and Aaron Vonk
Laboratory for Integrative Studies in Amphibian Biology, Group in Endocrinology, Museum of Vertebrate Zoology, and Dept. of Integrative Biology, University of California, Berkeley, CA 94720-3140 Address correspondence to: Tyrone B. Hayes, PhD Assoc. Professor Laboratory for Integrative Studies in Amphibian Biology Dept. of Integrative Biology University of California Berkeley, CA 94720-3104 Of 1-510-643-1054 Lab 1-510-643-1055 Fax 1-510-643-6264 Email firstname.lastname@example.org
- Environmental Health Perspectives http://ehp.niehs.nih.gov/
Key Words: atrazine, amphibian, endocrine disruption, hermaphrodite
Abbreviations: CA = California Co = company EPA = Environmental Protection Agency kg = kilogram km = kilometer NE = Nebraska ppb = parts per billion TX = Texas U.S. = United States
Atrazine is the most commonly used herbicide in the U.S. and probably the world. Atrazine contamination is widespread and can be present in excess of 1.0 part per billion (ppb) even in precipitation and in areas where it is not used. In the current study, we showed that atrazine exposure (= 0.1 ppb) resulted in retarded gonadal development (gonadal dysgenesis) and testicular oogenesis (hermaphroditism) in leopard frogs (Rana pipiens). Slower developing males even experienced oocyte growth (vitellogenesis). Furthermore, we observed gonadal dysgenesis and hermaphroditism in animals collected from atrazine-contaminated sites across the U.S. These coordinated laboratory and field studies revealed the potential biological impact of atrazine contamination in the environment. Combined with reported similar effects in Xenopus laevis, the current data raise concern about the effects of atrazine on amphibians in general and the potential role of atrazine and other endocrine disrupting pesticides in amphibian declines.
Atrazine is probably the most widely used herbicide in the world and one of the most common contaminants in ground and surface waters (US EPA 1994). Recently, Tevera- Mendoza et al. showed that atrazine-exposure (21 ppb) for as little as 48 hours resulted in severe gonadal dysgenesis in African clawed frogs (Xenopus laevis) (Tevera –Mendoza et al. 2002). Further, we showed that atrazine induced hermaphroditism at concentrations of only 0.1 ppb (Hayes et al. 2002) when administered throughout larval development.
Most water sources in the US, including rainwater, can exceed the effective concentrations in these laboratory studies (Hayes et al. 2002). Also, the concentration in our previous study (Hayes et al. 2002) is 30 times lower than the current drinking water standard (Hayes 1993). Despite the significance of the reported effects in X. laevis, both studies (Tevera-Mendoza et al. 2002; Hayes et al. 2002) were conducted in the laboratory on a single species. Whether the effects of atrazine are widespread in amphibians and whether effects occur in the wild remained unanswered.
In the current study, we examined the effects of atrazine on leopard frogs (Rana pipiens), a U.S. native species, under controlled laboratory conditions. Once effects of atrazine were identified, we examined wild R. pipiens from a variety of habitats in areas with reportedly low atrazine-use and areas with high atrazine-use in a transect that extended from Utah to the Iowa-Illinois border. Further, we collected water samples and examined atrazine contaminant levels at each site. These coordinated laboratory and field analyses uniquely addressed the ecological significance and relevance of our initial laboratory studies.
Materials and Methods
Animal care for laboratory studies. Leopard frogs (Rana pipiens) were obtained from Sensiba Marsh, Brown County, Wisconsin and shipped over-night to UCB. Eggs were allowed to hatch and then apportioned into rearing tanks. Larvae (30 per tank) were reared in 4 L of aerated 10% Holtfreter’s solution (Holtfreter 1931) and fed Purina rabbit chow (Purina Mills, St. Louis MO). Food levels were adjusted as larvae grew to maximize growth. Experiments were carried out at 22-23°C with animals under a 12-hour light-12 hour dark cycle (lights on at 6 a.m.).
Larval laboratory exposures. Larvae were treated by immersion with nominal concentrations of 0, 0.1, or 25 ppb atrazine (98% pure, Chemservice, Chester, PA). Concentrations were confirmed by chemical analysis (see below). Atrazine was predissolved in ethanol and all treatments received 0.0036% ethanol. Each treatment was replicated three times (30 larvae per replicate). Cages were cleaned, water changed, and treatments renewed every three days. All treatments were systematically relocated every three days to ensure that no treatments or tanks experienced position-effects. Animals were exposed throughout the larval period from two days post-hatching until complete tail reabsorption. In all experiments, all dosing and analyses were conducted blindly with color-coded tanks and treatments.
At metamorphosis (complete tail reabsorption), each animal was weighed and measured. Animals were euthanized in 0.2 % benzocaine (Sigma Chemicals, St. Louis MO), assigned a unique identification number, fixed in Bouin’s fixative and preserved in 70% ethanol until further analysis.
Histological analysis of gonads. All analysis was conducted blindly. Initially, the sex of all individuals was determined based on gross gonadal morphology using a Nikon SMZ 10A dissecting scope. In the laboratory study, histological analysis was conducted on nine females per treatment and on all males. All histology was conducted according to Hayes, ’95 (Hayes 1995). In brief, tissues were dissected and dehydrated in graded alcohols followed by infiltration with histoclear and paraffin. Serial istological sections were cut at 8 µ through the entire gonad. Slides were stained in Mallory’s trichrome stain and analyzed using a Nikon Optiphot 2 microscope. Images of gonads were recorded using a Sony DKC-5000 digital camera. For gonadal analysis, we examined every section from each gonad.
Site-selection for field studies. Initially, we chose study localities based on atrazine-use, as determined by atrazine sales (Figure 1). All localities were between 39° N and 43° N latitude. Counties with less than 0.4 kg/km2 atrazine-use were chosen as potential control sites and areas with > 9.3 kg/km2 atrazine-use were chosen as potential atrazine-exposed sites. We began sampling in Utah on July 15, 2001 and moved eastward. In Utah, we chose one site (Juab County) in an area with less than 0.4 kg/km2 atrazine-sales, and we collected in Cache County with 0.4-2.4 kg/km2 reported atrazine-use. Carbon County, Wyoming, was considered a control site, because the locality is not in the vicinity of farms and the county (most of the state, in fact) reports less than 0.4 kg/km2 atrazinesales. In Nebraska, we chose one site in York County with high atrazine-use, and one site in Cherry County where atrazine sales were below 0.4 kg/km2. All sites in Iowa were considered exposed sites, except a single site in a wildlife protection area in Iowa. We stopped sampling at the Iowa-Illinois border, because Rana pipiens populations are reportedly low or threatened in Illinois and Indiana.
Frog and water sampling from field localities. At each site (Figure 1), we collected 100 animals (eight sites, for a total of 800 animals). We selected small individuals in an attempt to sample newly metamorphosed animals. Immediately after collection, animals were euthanized in benzocaine, fixed in Bouin’s fixative for 48 hours, and preserved in 70% ethanol. Animals were returned to the laboratory, measured, the sex of each determined, and histological analysis conducted on the gonads of 20 males from each site and a subset of females from each site. Histology was conducted as described for laboratory studies.
At each site, we collected water (100 ml) in clean chemical-free glass jars (Fisher Scientific Co, Houston, TX) for chemical analysis. Water samples were frozen on dry ice immediately upon collection and maintained frozen (-20°C) until analysis. Atrazine levels from all sites were determined by PTRL West Inc. (Hercules, CA). Water samples were extracted in organic solvent, followed by aqueous/organic extraction. Samples were analyzed by liquid chromatography/ mass spectrophotometry using the daughter ion. Duplicate samples were analyzed at the Hygienic Laboratory (University of Iowa, Iowa City, Iowa). The Hygienic Laboratory used EPA method 507 (U S EPA). In brief, water samples were extracted in organic solvent and subjected to gas chromatography with a nitrogen phosphorous detector. Both analytical laboratories received coded samples and were not aware of the collection localities. In addition each laboratory received negative controls that contained only Holtfreter’s solution and positive controls that contained mixtures of pesticides at both 0.1 ppb and 10 ppb.
Detection limits were 0.1 ppb for both laboratories and data for duplicated samples were accepted only when both laboratories reported results within 10% of each other. Reported values reflect the higher estimate rounded to the nearest 0.1. In addition to atrazine, PTRL West Inc. reported the atrazine metabolites, diaminochlorotriazine (DAC), deisopropylatrazine (DIA), deethylatrazine (DEA) as well as the triazines (simazine and hexazinone) and two other herbicides (diuron, and norflurazon) from all sites. In addition to atrazine, the following pesticides were reportedly used at Site 5 in Nebraska: herbicides (metolachlor, alachlor, glyphosate), fungicides (metalaxyl, nicosulfuron, propiconazole), insecticides (ß-cyfluthrin, ?-cyhalothrin, tebupirimphos) and were analyzed using appropriate methods by the Iowa Hygienic laboratories for site 1 (UT), site 3 (WY), and Site 5 (NE) only.
Gonadal analysis in laboratory-reared animals. Control animals were sexually differentiated at metamorphosis. The earliest males to metamorphose had solid testicular lobules. Primary spermatogonia were recognized in the lobules of some animals (Figure 2). Animals that metamorphosed later had open lobules that contained both primary spermatogonia and spermatids. Females had numerous oocytes in their gonads and a central ovarian cavity (Figure 3).
Atrazine-treated males (0.1 and 25 ppb) were sexually differentiated at metamorphosis as well, but 36% and 12% of the males treated with 0.1 and 25 ppb atrazine, respectively, suffered from gonadal dysgenesis--under-developed testes with poorly structured, closed lobules (or no lobules at all) and low to absent germ cells (Figure 4). Further, 29% of the 0.1 ppb-treated animals and 8% of the animals treated with 25 ppb displayed varying degrees of sex-reversal. The testicular lobules of sexreversed males contained oocytes when observed histologically (Figure 5) and males that metamorphosed later contained large numbers of oocytes (Figures 6-7). Males that appeared to undergo complete sex-reversal had gonads almost completely filled with oocytes and only a limited number of lobules remained (Figure 7). In two males, oocytes were vitellogenic and protruded through the testicular lobules, which made the oocytes observable upon gross analysis of the gonads (Figure 8). Control males never contained testicular oocytes, although two control males contained two-three degenerating extragonadal oocytes (not within lobules) and a single control male showed gonadal dysgenesis (Fig, 9). There were no observable effects in atrazine-treated females.
Once effects were identified in laboratory-reared animals, we conducted a study of gonadal morphology in field-collected R. pipiens to determine if animals exposed in the wild displayed similar abnormalities. Localities for collections are shown in Figure 1. We chose four sites from potential control/uncontaminated areas (counties in Utah, Wisconsin, and Nebraska that reported atrazine sales below 0.4 kg/km2 and a nonagricultural site in Iowa) and four contaminated areas (Cache County, Utah, the single county that reported more than 0.4 kg/km2 in atrazine sales, one site in an agricultural area in Nebraska and two similar sites in Iowa). In addition to varying in the amount of atrazine-use, the habitats at collecting sites ranged from wildlife protection areas to agricultural runoff and cornfields (Fig. 10, Table 1). Chemical analysis of water samples collected from each site revealed that none of the sites were atrazine-free, and only one site (Juab County, Utah) had atrazine levels below 0.2 ppb (Table 1). Sites in Utah and Wyoming did not have detectable levels of atrazine metabolites (Fig. 11a). None of the other pesticides assayed were present at any site except metolachlor, which was present at site 5 (York, NE) at 0.39 ppb.
Analysis of gonads from field-collected animals. Testicular oocytes were identified in males from seven of eight sites (Table 1, Fig. 11b). All sites associated with atrazine sales that exceeded 0.4 kg/km2 and atrazine contaminant levels that exceeded 0.2 ppb had males that displayed sex-reversal similar to those abnormalities induced by atrazine in the laboratory (Figs. 12-15). Also, in high-use York County, Nebraska, 28% of the males examined had gonadal dysgenesis (Figure 13) and testicular oocytes were found in a single male. The poorly developed testicular lobules that lacked germ cells observed in males from the corn field in York County resembled gonadal dysgenesis observed in males exposed to 0.1 ppb atrazine in our laboratory study and effects described in Xenopus laevis (Tevera-Mendoza et al. 2002). Site 3, on the North Platte River in Wyoming, was not associated with direct agricultural runoff, but had the highest incidence and the most advanced cases of hermaphroditism. Most of the males observed from this site (92%) had testicular oocytes and many animals showed advanced stages of complete sex-reversal. All other sites had varying frequencies and severities of gonadal abnormalities. There were no observable abnormalities in females from any of the localities.
Atrazine-exposure disrupted gonadal development in exposed larvae. Testicular tubules were poorly developed in exposed animals (gonadal dysgenesis), germ cells appeared reduced, and oocytes were allowed to develop (testicular oogenesis) in animals identified as hermaphrodites. In at least two animals, oocytes were vitellogenic and protruded through the testes. Furthermore, effects were more pronounced at the lower dose (0.1 ppb). Widespread atrazine contamination was accompanied by observations of hermaphroditic animals in the field. Combined, these studies suggest that atrazine impacts amphibians in the wild.
Relevance to previous work. A previous study examined the effects of atrazine on Rana pipiens (Allran and Karasov 2000), but used much higher doses and did not examine gonadal differentiation. Thus, this previous study did not identify the abnormalities that we observed here. The current study supports our previous findings of atrazine-induced hermaphroditism in Xenopus laevis (Hayes et al. 2002) as well as the findings of Tevera- Mendoza et al. (2002), who showed retarded gonadogenesis and decreased germ cell numbers in atrazine-exposed males. Males with testicular oogenesis had lobed gonads similar to gonads of some atrazine-exposed males described in our studies of Xenopus laevis. Furthermore, atrazine-exposure resulted in testicular oogenesis and even induced growth (vitellogenesis) of the oocytes in slower developing males, but had no effect in females. Atrazine does not bind the estrogen receptor (Tennant et al. 1994), but studies in reptiles (Crain et al. 1997), mammals (Sanderson et al. 2000; Sanderson et al. 2001), and fish (Sanderson et al. 2001) showed that atrazine induces aromatase and thereby increases the production of endogenous estrogen. The demasculinization (failure to induce spermatogenesis) and feminization (induction and growth of oocytes) observed in the current study and previous work (Tevera-Mendoza et al. 2002; Hayes et al. 2002) are explainable via the proposed mechanism.
Effects on germ cell differentiation. Witschi (1929) suggested that some species/populations of Rana display “rudimentary hermaphroditism” in which “undifferentiated” races of Ranid frogs had ovaries with eggs anteriorally and testicular nodules (which did not contain oocytes) posteriorally. This developmental pattern was not observed in the population of Rana pipiens used in our current study. Even atrazinetreated animals did not display the morphology described by Witschi. In atrazine-exposed male, testicular oocytes were always in the posterior section of the gonads. Also, testicular oogenesis and hermaphroditism were never observed in control animals in the current study or in other unpublished observations of R. pipiens in our laboratory including more than 7,000 individuals and four populations (Utah, Nebraska, Wisconsin, and Connecticut).
Three control males contained up to three extragonadal degenerating oocytes (never within the testicular lobules) at the posterior end of the gonads, however. Normally, germ cells migrate into the developing gonad from the yolk or gut endoderm (depending on the species). Primordial germ cells that fail to enter the developing testes become oocytes by default, even in mammals (McClaren 1995; Nakatsuji and Chuma 2001), but eventually degenerate. The current data suggest that atrazine demasculinized the gonads of exposed males. Instead of releasing the putative spermatogenesis-inducing factor (Witschi 1957), which would result in the degeneration of oocytes, atrazineexposed males supported differentiation and even growth of these oocytes.
The observation that 0.1 ppb atrazine was more effective than the higher dose (25 ppb) is interesting. Both the proportion of males with gonadal dysgenesis and the proportion with testicular oocytes (hermaphrodites) were higher at the lower dose. Low- dose effects have been described for a number of endocrine disruptors (estrogenic compounds and anti-androgenic compounds) and some compounds even produce different effects at different doses and in different tissues (Akingbemi and Hardy 2001). Low-dose effects of demasculinizing and feminizing environmental endocrine disruptors on male development have been of particular concern (Akingbemi and Hardy 2001). Furthermore, similar perplexing effects are known for estradiol 17ß on sex differentiation in Rana pipiens (Richards and Nace 1978). Low doses of estradiol (< 0.07 µM) do not affect sex differentiation, higher doses (0.07 – 0.18 µM) produce 100% females, and still higher doses (> 3.69 µM) produce 100% males (Hayes 1998). A mixture of normal males, females, and intersexes are produced at doses between 0.18 and 3.69 µM.
Relevance to wild amphibian populations. The use of a U.S. native species in the current study allowed us to assess the realized impact of atrazine on wild amphibians. Wild populations that contained males with gonadal dysgenesis, and testicular oocytes (hermaphrodites) were associated with localities with atrazine use and/or atrazine contamination. Reeder et al. (1998) described testicular oocytes in field-collected frogs (Acris creptians) and suggested that atrazine may be involved in this abnormality, but did not have laboratory data to support the suggestion. Atrazine may not be the only compound that induces testicular oogenesis, however. There may be many chemicals, natural products, and even populations that naturally display this phenomenon (Witschi 1929). Nevertheless, the current study showed that atrazine induced testicular oogenesis and hermaphroditism in a population that does not show this developmental pattern under controlled laboratory conditions and that hermaphroditism in wild R. pipiens is associated with atrazine-use and contamination.
Extent of atrazine contamination. Our current study demonstrated the extent of atrazinecontamination and its potential impact. The locality in Wyoming (North Platte River) with the highest frequency of sex-reversal (92% of the males) is not in the vicinity of farms, nor is it in a county that reports significant atrazine-use. The North Platte River is fed by streams from Colorado. Atrazine contamination of the Platte River Valley in Colorado is well-documented and the contamination flows into Wyoming via the Platte River system (Kimbrough and Litke 1995). Thus, amphibians in Wyoming, at a locality that does not report significant atrazine-use, are at risk from contamination from Colorado. Similarly, atrazine contamination of ground and well water has been reported in Utah, even in areas where atrazine is not used heavily (Thiros 2000). Contamination of ground water in Utah, transport of atrazine to Wyoming via the North Platte River, the presence of atrazine in Cherry County Nebraska (where no atrazine-use is reported) and findings of hermaphrodites at these localities further demonstrate the problem of widespread atrazine-contamination.
Additional water collections and contaminant analysis revealed the difficulties in determining the contaminant levels that larvae experience. Even though atrazine reportedly has a short half-life (as little as eight days) (Solomon et al. 1996), we measured atrazine-contamination (> 0.2 ppb) at irrigation ditches in York County, Nebraska, even on March 31, 2001. According to on-site pesticide application records at the time of water collections, atrazine had not been applied since May 19, 2000. Thus, atrazine levels in this area never decreased below the determined biologically active levels. Furthermore, atrazine levels varied from 15.2 ppb to 0.8 ppb over a 24-hour period (July 22-23, 2001, respectively) as a result of evaporation followed by an increase in the water level from irrigation and runoff. Also, even though atrazine is applied directly only twice per year at this locality, the water source used for irrigation had atrazine levels of 0.7 ppb, so levels of atrazine capable of inducing testicular oogenesis are continuously applied to these fields. In addition, runoff from cornfields in this area flowed into adjacent organic farms and wildlife protection areas, resulting in atrazine contamination in excess of 15 ppb at both the organic farm and in the refuge on July 23, 2001 even though atrazine had not been applied since May 13, 2001 at this locality. Further, on July 23, 2001, atrazine levels in rainwater and tap water in York County were 0.4 ppb and 0.3 ppb, respectively. Thus, even rain and tap water in York, NE contained enough atrazine to disrupt normal male development in amphibians. Finally, as suggested in Figure 16, atrazine levels were likely at their lowest at the time of our collections in July and the levels likely peak during critical stages of larval development.
Difficulties with quantitative analyses and predictive capabilities. Although we attempted to predict localities where hermaphrodites might occur based on atrazine-use, the movement of atrazine into areas such as Carbon County, Wyoming via the North Platte River, the presence of atrazine in Cherry County, Nebraska (where atrazine use is not reported), and local use of atrazine in areas that do not report high-use, make such predictions difficult. In addition, habitat type and local land-use history are not good predictors because of the transport of atrazine, e.g. the hermaphrodites identified in the wildlife area in Iowa, Cherry County, NE, and Carbon County, WY.
Further, low-dose effects and the extent and variability in atrazine-contamination make quantitative analyses difficult. Because 0.1 ppb was effective in the laboratory (and, in fact, more effective than 25 ppb) we did not identify a concentration that is below threshold in our laboratory studies. In X. laevis (Hayes et al. 2002), 0.01 ppb was ineffective but hermaphroditism was observed at 0.1 – 200 ppb. Even if a minimum concentration were identified, both the current and the previous study (Hayes et al. 2002) suggest that there is not a linear-dose response. In fact, both studies imply that there is an “inverted U” (parabolic) response (Chen 2001) in which very low concentrations may be without effect, higher concentrations have decreasing effectiveness, and intermediate low concentrations are most effective. There does not appear to be a relationship between atrazine concentrations and the number or size of testicular oocytes, but longer exposuretimes may be associated with increasing numbers of testicular oocytes, size of testicular oocytes, and the extent of testicular conversion to ovaries.
Even if the dose-response pattern were understood, variation in atrazine levels from locality to locality and even from day to day at a single locality make it difficult to predict where high frequencies of affected males might occur. In addition such statistical models would involve non-parametric statistics (such as G-tests) that rely on testing observed frequencies of hermaphrodites against predicted frequencies. Either expected frequencies would be set to zero, or we would assume some natural expected frequency of hermaphroditism that may vary between populations. It is difficult to determine the predicted (natural) frequencies, although we have reared animals (more than 7,000) from Utah, Nebraska, Wisconsin, and Connecticut in the laboratory and never observed testicular oogenesis or hermaphroditism unless animals were exposed to atrazine. Despite difficulties that limit quantitative analyses at this time, testicular oogenesis and hermaphroditism always occurred at localities associated with atrazine-exposure and were absent only at the single site with < 0.2 ppb atrazine contamination (Juab County, Utah).
The threat to amphibians. Findings of similar effects of atrazine on sexual development in two diverse species (Xenopus laevis and Rana pipiens), show that effects of atrazine are not restricted to a single species and, in fact, likely present a problem for amphibians in general. The pattern of atrazine-use creates even more concern. As shown in Figure 16, atrazine levels are highest during larval development (Conant 1998; Stebbins 1985) for most local species. Applied as a pre-emergent, atrazine contamination of water sources peaks with spring rains. The timing of atrazine contamination of water sources directly coincides with amphibian breeding activities, as many amphibians reproduce during early spring rains. Given the identified effects of atrazine in the laboratory, combined with the apparent correlation of atrazine-contamination with similar morphologies in the wild and the pattern of atrazine-use, the potential impact of atrazine on amphibians is significant. Many amphibian species are in decline (Wake 1991; Blaustein and Kiesecker 2002; Gardner 2001), and Rana pipiens populations are declining in many locations in Indiana and Illinois. Juvenile Rana pipiens were abundant at all of our collection sites, however, including agricultural areas in Iowa and Nebraska. The abundance of frogs at these sites suggests that the effects are reversible, that some percentage of the population does not show this response, that these developmental abnormalities do not impair reproductive function at sexual maturity, and/or that continuously exposed populations have evolved resistance to atrazine. In fact, although gonadal dysgenesis may be induced by atrazine (based on our laboratory studies), it may be a mechanism of resistance as well. If lobular formation and germ cell differentiation are delayed until after metamorphosis, then portions of the population that display gonadal dysgenesis may escape atrazine-induced sex-reversal, because they would undergo sex differentiation after metamorphosis once they leave the contaminated water. This hypothesis is testable in the laboratory, as the proportion of exposed males with gonadal dysgenesis at metamorphosis should reflect the proportion of normal males in the population after metamorphosis. In addition, higher proportions of affected males at a locality that only periodically experience high contaminant levels (such as Wyoming) may reflect that these populations have not been under the same intensity of selection for atrazine resistance. Periodic fluctuations in atrazine contamination may affect large proportions of some populations from year to year, but unexposed animals, or animals from previous years may continue to breed. Further studies are needed to address these questions and the realized impact of atrazine on exposed populations.
There are likely many factors involved in amphibian declines. Endocrine disruption by pesticides is but one potential cause and atrazine only one such compound. However, given the widespread use and ubiquitous contamination by atrazine, its pattern of use, and its potency as an endocrine disruptor, atrazine likely has a significant impact on amphibian populations. In particular, given recent evidence that atrazine potentiates parasitic infections in amphibians (Kiesecker 2002) in addition to its impact on reproductive development, the role of atrazine in amphibian declines is of particular concern. Further, enhancement of atrazine effects when mixed with other pesticides, as indicated in our ongoing studies, must be explored.
Akingbemi BT, Hardy MP. 2001. Oestrogenic and antiandrogenic chemicals in the environment: Effects on male reproductive health. Ann Med 33:391-403.
Allran JW, Karasov WH. 2000. Effects of atrazine and nitrate on northern leopard frog (Rana pipiens) larvae exposed in the laboratory from posthatch through metamorphosis. Environ Toxicol Chem 19:2850-2855.
Battaglin WA, Goolsby DA. 1995. Spatial data in geographic information system format on agricultural chemical use, land use, and cropping practies in the United States. U.S. Geological Survey Water-Resources Investigations Report. 94-4176.
Blaustein AR, Kiesecker, JM. 2002. Complexity in conservation: Lessons from the global decline of amphibian populations. Ecol Lett 5(4):597-608.
Chen CW. 2001. Assessment of endocrine disruptors: Approaches, issues, and uncertainties. Folia Histochem Cytobiol 39(suppl 2):20-23.
Conant RA. 1998. Field Guide to Reptiles and Amphibians: Eastern and Central North America. Boston:Houghton Mifflin.
Crain DA, Guillette LJ Jr, Rooney AA, Pickford DB. 1997. Alterations in steroidogenesis in alligators (Alligator mississippiensis) exposed naturally and experimentally to environmental contaminants. Environ Health Perspect 105:528-533.
Gardner T. 2001. Declining amphibian populations: a global phenomenon in conservation biology. Animal Biodiversity and Conservation 24(2):25-44.
Goolsby DA, Pereira WE. 1995. Pesticides in the Mississippi River. In: Contaminants in the Mississippi River, 1987-92 (Meade RH, ed). U.S. Geological Survey Circular 1133, 87-102.
Hayes E. 1993. EPA’s chemical information data base (Keith LH, ed). EPA Journal Jan./Feb./Mar., 48.
Hayes TB, et al. In press. Hermaphrodites beyond the cornfield: Atrazine-induced testicular oogenesis in leopard frogs (Rana pipiens). Nature.
Hayes TB, Collins A, Lee M, Mendoza M, Noriega N, Stuart AA, Vonk A. 2002. Hermaphroditic, demasculinized frogs after exposure to the herbicide atrazine at low ecologically relevant doses. Proc Natl Acad Sci U S A 99:5476-5480.
Hayes TB. 1998. Sex determination and primary sex differentiation in amphibians. J Exp Zool 281:373-399.
Hayes TB. 1995. An histological examination of the effects of corticosterone in larvae of the Western Toad, Bufo boreas (Anura: Bufonidae), and the Oriental Fire-bellied Toad, Bombina orientalis (Anura: Discoglossidae). J Morphol 226:297-307.
Holtfreter J. 1931. Uber die Aufzucht isolierter Teile des Amphibian Keimes II. Arch F Ent Mech 124:404-465.
Kiesecker JM. 2002. Synergism between trematode infection and pesticide exposure: a link to amphibian deformities in nature. Proc Natl Acad Sci U S A. 99:9900-9904.
Kimbrough RA, Litke DW. 1995. Pesticides in surface water in agricultural and urban areas of the South Platte River Basin from Denver, Colorado to North Platte, Nebraska, 1993-94. NAWQA Program. South Platte River Basin Study.
McClaren A .1995. Germ cells and germ cell sex. Phil Trans R Soc Lond B 350:229-233.
Nakatsuji N, Chuma S. 2001. Differentiation of mouse primordial germ cells into female or males germ cells. J Dev Biol 45:541-548.
Reeder AL, Foley GL, Nichols DK, Hansen LG, Wikoff B, Faeh S, et al. 1998. Forms and prevalence of intersexuality and effects of environmental contaminants on sexuality in cricket frogs (Acris crepitans). Environ Health Perspect 106:261-266.
Richards CM, Nace GW. 1978. Gynogenetic and hormonal sex reversal used in tests of the XX-XY hypothesis of sex determination in Rana pipiens. Growth 42:319-331.
Sanderson JT, Letcher RJ, Heneweer M, Giesy JP, van den Berg M. 2001. Effects of chloro-s-triazine herbicides and metabolites on aromatase activity in various human cell lines and on vitellogenin production in male carp hepatocytes. Environ Health Perspect 109:1027-1031.
Sanderson JT, Seinen W, Giesy JP, van den Berg M. 2000. 2-chloro-triazine herbicides induce aromatase (CYP19) activity in H295R human adrenocortical carcinoma cells: a novel mechanism for estrogenicity? Toxicol Sci 54:121-127.
Solomon K, Keith R, David B, Baker R, Richards P, Dixon KR, et al. 1996. Ecological risk assessment of atrazine in North American surface waters. Environ Toxicol Chem 15:31-76.
Stebbins R. 1985. A Field Guide to Western Reptiles and Amphibians: Field Marks of all Species in Western North America, Including Baja California. Boston:Houghton Mifflin.
Tennant MK, Hill DS, Eldridge JC, Wetzel LT, Breckenridge CB, Stevens JT. 1994. Chloro-s-triazine antagonism of estrogen action: Limited interaction with estrogen receptor binding. J Toxicol Environ Health 43:197-211.
Tevera-Mendoza L, Ruby S, Brousseau P, Fournier M, Cyr D, Marcogliese D. 2002. Response of the amphibian tadpole (Xenopus laevis) to atrazine during sexual differentiation of the testis. Environ Toxicol Chem 21:527-531.
Thiros SA. 2000. Quality of shallow ground water in areas of recent residential and commercial development in Salt Lake Valley, Utah 1999. USGS Fact Sheet 106-00.
U.S. Environmental Protection Agency. 1994. Atrazine, simazine, and cyanizine. Notice of initiation of special review. Fed Reg 59:60412-60443.
U. S. Environmental Protection Agency. Methods for Determination of Organic Compounds in Drinking Water. EPA/ 600/R-95-131.
Wake DB. 1991. Declining amphibian populations. Science 253:860.
Witschi E. 1957. The inductor theory of sex differentiation. J Fac Sci Hokkaido Univ Ser VI 13:428-439.
Witschi E. 1929. Rudimentary hermaphroditism and Y chromosome in Rana temporaria. J Exp Zool 54:157-223.
We thank Alton Jones Foundation, World Wildlife Fund, Homeland Foundation, and the Rose Foundation for funding the current work. The Howard Hughes Biology Scholars/Fellows Program funded Haston and Tsui. Adrian Brunner-Brown assisted with field collections and Paola Case assisted in laboratory analysis. We thank Dr. Luis Ruzo for assistance with contaminant analysis. Krissie Wilson provided assistance in fieldwork in UT, Mike Fritz in NE, and Fred Janzen in IA. Kathleen LeVering provided eggs from Wisconsin. Tom Borden and Paul Aldridge documented field-work. Several farm owners allowed us to work on their property in NE and IA. Also, we thank Katherine Kim for her support.
Table 1. Dates, Localities, descriptions, and atrazine levels for field localities.
Site Date State County Latitude Longitude Alt1 Source Habitat Atrazine (ppb) 1 07/15/01 UT Juab 111°52.23W 39°46.63N 1500 Pond Graze land 0.1 2 07/17/01 UT Cache 111°50.14W 39°43.40N 1555 Pond Golf course 0.2 3 07/19/01 WY Carbon 107°03.28W 41°51.68N 1952 River Wildlife area 0.2 4 07/23/01 NE Cherry 101°42.89W 42°41.67N 1031 Pond Prairie 0.3 5 07/22/01 NE York 97°22.38W 40°55.88N 480 Ditch Corn field 0.8 6 07/26/01 IA Polk 93°27.39W 41°48.11N 252 Ditch Corn field 6.7 7 07/28/01 IA Polk 93°25.50W 41°47.47N 246 Marsh Wildlife area NA2 8 07/28/01 IA Clinton 90°21.28W 41°44.46N 211 Stream River valley 0.5 1 Alt = altitude (in meters) 2 NA = not available. Levels reported by the two analytical laboratories were inconsistent (not within 10%). One laboratory (PTRL West) reported non-detectable levels.
Figure 1. Map of the United States showing atrazine-use based on sales (Battaglin and Goolsby 1995). The pink overlay shows the natural range for leopard frogs (Rana pipiens) in the U.S. Numbers indicate sites where water (for chemical analysis) and frogs (for histological analysis of gonads) were collected. Collection sites are numbered and correspond to site numbers used in Table 1. Reprinted with permission from Hayes et al. (in press)
Figures 2-3 (not included) Gonadal morphology and histology of a control male and female. The yellow coloration (in 2a and 3a) is due to fixation in Bouin’s solution. Histological crosssections were 8 µ thick and slides were stained in Mallory's trichrome stain. Bar (in panel 3b) = 1.0 mm for panels 2a and 3a and 250 µ for 2b,c and 3b. Figure 2 shows the right testis of a control male leopard frog (Rana pipiens) at metamorphosis. Panel 2a, shows the gonad still attached to the kidney. White arrows show the position of transverse crosssections shown in Panel 2b (anterior section) and Panel 2c (posterior section). Testicular lobules were well developed and the specimen shown had both primary spermatogonia and spermatids present. In general, testes were more differentiated (contained more distinct tubules and germ cells) anteriorally than posteriorally. R = rete testis. Figure 3 shows the right ovary from a control female. Panel 3a, shows the gonad still attached to the kidney. The white arrow shows the position from which the transverse cross-section shown in panel 3b was taken. The ovary contains a large number of oocytes. The ovarian cavity is visible.
Figures. 4-8 (not included) Testes from males treated with 0.1 ppb atrazine. The animals shown experienced either gonadal dysgenesis (Figs. 4a-b) or testicular oogenesis and hermaphroditism (Figs. 5-8). The yellow color of the gonads (in a) is due to fixation in Bouin's solution. All histological cross-sections were 8 µ thick and slides were stained in Mallory's trichrome stain. Bar (in panel 4a) = 250 µ for photomicrographs of gonads (panel a for all figures). Figure 4 shows the testes of a male with gonadal dysgenesis. Panel 4b shows a transverse cross-section through the area indicated by the arrow in Panel 4a. Histological analysis revealed that the individual had poor lobular development and the gonads were devoid of germ cells. Bar (in 4b) = 100 µ. Figure 5 shows the gonad of an animal with lobed gonads. Arrows (in panel 5a) show areas where transverse cross-sections were taken. Panel 5b shows the anterior-most section. The testicular lobules are open and distinct and contain spermatids. Panel 5c shows the center section, which contained three lobules. Panel 5d shows the most posterior section. Welldeveloped lobules contain mostly spermatids, but one lobule is devoid of spermatids or spermatogonia and contain single large oocytes each. A single oocyte is seen in transverse cross-section. Bar = 100 µ. Note that the scale for panels 4b and panels 5b-d are the same. Figure 6 shows the right lobed gonad of a male with anterior testes and developing posterior ovary. White arrows show areas where transverse cross-sections were taken. Panels 6b and c show transverse cross-sections from anterior to posterior. The anterior portion is testicular with lobules that contain spermatids. White arrows in 6b indicate lobules with spermatids. The posterior portion of the testis (6c) has large testicular oocytes. Bar (in 6c) = 250 µ for panels 6b-c. Figure 7 shows the left testis of the same animal shown in Figure 6. Panel 7b shows a sagittal section of the left testis, which is almost completely converted into an ovary. The entire posterior gonad is ovarian and oocytes are beginning to grow in the anterior portion. A small portion of the gonad (outlined by the white box in 7b and magnified in panel 7c) still contains testicular lobules, but the lobules lack germ cells. Bar = 250 µ for panel 7b. Figure 8 shows gonads from a male with vitellogenic testicular oocytes. The posterior portion of the gonad is filled with oocytes that are protruding through the testicular lobules and can be seen on the surface of the gonad (8a). Transverse cross-sections indicated by the white arrows show that the anterior testis has poorly developed testicular lobules (8b). The black arrowhead in 8b shows a tangentially sectioned oocyte. Panels 8c-d show large vitellogenic oocytes in the posterior portion of the gonads. Bar (in 8d) = 250 µ for panels 8b-d.
Figure 9. (not included) Frequency (percent) of gonadal abnormalities in males treated with atrazine in the laboratory.
Figure 10. Habitats at collection localities where animals and water were collected for analysis: Site 1, Juab County, UT (a), Site 2, Cache County, UT (b), Site 3, Carbon County, WY (c), Site 4, Cherry County, NE (d), Site 5, York County, NE (e), Site 6, Polk County, IA (f), Site 7, Polk County, IA (g), and Site 8, Clinton County, IA (h) Detailed coordinates and descriptions of habitats are given in Table 1. Figure 11. Presences of atrazine and metabolites: atrazine (ATZ), deisopropyl atrazine (DIA), deethylatrazine (DEA), and Diaminochloroatrazine (DAC) at Sites 1-8 (top) and frequency (percent) of gonadal abnormalities in males collected from the wild (bottom). Localities and descriptions of each site are found in Table 1. Atrazine contaminant levels for each site are also indicated by the stars on the bottom figure. Bottom figure reprinted with permission (Hayes et al. in press).
Figure 12-15 (not included) Gonads of males collected from various localities showing gonadal abnormalities of varying severities. The yellow color in photographs of gonads (a) is due to fixation in Bouin's solution. All histological cross-sections were 8 µ thick and slides were stained in Mallory's trichrome stain. Bar (in 13a) = 0.1 mm for photomicrographs of gonads. Figure 12 shows a male from site 8 (Clinton, IA). Panel 12a shows the left testis. The white arrow in 12a shows the area where the transverse cross-section was taken. Panel 12b shows well-developed testicular lobules with spermatids and three lobules that contain both spermatids and a single large oocyte each. Bar (in 12b) = 250 µ. Figure 13 shows an animal from site 5 (York County, NE). The animal shown displayed gonadal dysgenesis as seen in 28% of the animals from that locality. Panel 13b and c show the anterior and posterior cross sections, respectively, taken from the areas indicated by the white arrows in 13a. This morphology was similar to that displayed by 36% of the animals exposed to 0.1 ppb atrazine in the laboratory. Bar (in 13b) = 250 µ. Figure 14 shows an animal from Carbon County, Wyoming. Panel 14b shows numerous testicular oocytes, in fact, filling 100% of the testicular lobules. The inset (panel 14c) is a magnified view of the boxed area in panel 14b, and shows that some lobules had as many as three oocytes. Bar (in 14b) = 500 µ. Figure 14b reprinted with permission (Hayes et al. in press). Figure 15 shows a hermaphrodite undergoing what appears to be complete sex-reversal. This specimen from Carbon County, Wyoming has gonads that are becoming convoluted similar to an ovary (Panel 15a). Panels 15b-c reveal that the gonads contain numerous oocytes. The animal has developed an ovarian cavity and lost its lobular structure. Bar (in 15c) = 250 µ.
Figure 16 Atrazine contamination along the Mississippi River from 1991 to 1992 (26 Goolsby and Pereira 1995). The horizontal axis shows months over the two years. The dashed horizontal line shows the maximum contaminant level set by the US EPA (3 ppb) and the red horizontal line shows the dose effective at producing hermaphrodites in the laboratory (0.1 ppb). The red vertical overlays for each year show the timing of larval development for amphibians in each region.
If you have come to this page from an outside location click here to get back to mindfully.org