![The relationship between time since predator (Pred) cue addition and exposure to atrazine (Atr) or solvent (Solv) on the mean location (AE1 standard error [SE] to reduce overlap of SE bars) of tadpoles. Positive values represent attraction to the predator cue, and negative values represent avoidance of the predator cue.](https://www.researchgate.net/profile/Jason-Rohr/publication/291516798/figure/fig1/AS:597735564644352@1519522672482/The-relationship-between-time-since-predator-Pred-cue-addition-and-exposure-to-atrazine_Q320.jpg)
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Environmental Toxicology
THE HERBICIDE ATRAZINE INDUCES HYPERACTIVITY AND COMPROMISES TADPOLE
DETECTION OF PREDATOR CHEMICAL CUES
MACKENZIE EHRSAM,SARAH A. KNUTIE, and JASON R. ROHR*
Department of Integrative Biology, University of South Florida, Tampa, Florida, USA
(Submitted 31 August 2015; Returned for Revision 9 November 2015; Accepted 20 January 2016)
Abstract: The ability to detect chemical cues is often critical for freshwater organisms to avoid predation and find food and mates. In
particular, reduced activity and avoidance of chemical cues signaling predation risk are generally adaptive behaviors that reduce prey
encounter rates with predators. The present study examined the effects of the common herbicide atrazine on the ability of Cuban tree frog
(Osteopilus septentrionalis) tadpoles to detect and respond to chemical cues from larval dragonfly(Libellulidae sp.) predators. Tadpoles
exposed to an estimated environmental concentration of atrazine (calculated using US Environmental Protection Agency software;
measured concentration, 178 mg/L) were significantly hyperactive relative to those exposed to solvent control. In addition, control
tadpoles significantly avoided predator chemical cues, but tadpoles exposed to atrazine did not. These results are consistent with previous
studies that have demonstrated that ecologically relevant concentrations of atrazine can induce hyperactivity and impair the olfactory
abilities of other freshwater vertebrates. The authors call for additional studies examining the role of chemical contaminants in disrupting
chemical communication and the quantification of subsequent impacts on the fitness and population dynamics of wildlife. Environ
Toxicol Chem 2016;9999:1–6. #2016 SETAC
Keywords: Amphibian Behavior Infodisruption Olfaction Predation
INTRODUCTION
Infodisruption, defined as chemical contaminants disrupting
communication within or among organisms, has 2 modes of
expression: 1) an external interorganismal disruption that causes
a breakdown in detection or production of chemical signals
between senders and receivers, and 2) an internal intra-
organismal disruption that can affect cell-to-cell communica-
tion [1]. Several chemical contaminants are infodisruptors, and
both internal and external infodisruption can affect organisms’
health [1,2]. For many organisms, external infodisruption can
interfere with locating predators, mates, and food [1,2].
Similarly, endocrine disruptors, a type of internal organismal
infodisruptor (which eventually can also affect communication
between the sexes) [1], have been implicated in the collapse
of wildlife populations [3]. Unlike internal infodisruptors
(e.g., endocrine disruptors), external infodisruptors that
affect communication among individuals have not been well
studied [1].
For freshwater organisms, chemical cues are often cru-
cial [4–6]. This is because visual and auditory cues tend to
attenuate rapidly through water. In addition, turbidity in
freshwater ecosystems can further make visual cues unreliable.
Consequently, most freshwater organisms rely heavily on
chemical communication to avoid predation and find food and
mates [7,8], which is why external interorganismal infodisrup-
tors have the potential to adversely affect the fitness of
freshwater organisms [1,2].
One of the most common chemical contaminants in
freshwater ecosystems is the widely used herbicide 2-chloro-
4-(ethylamino)-6-(isopropylamino)-S-triazine (atrazine) [9].
Because of its heavy use, persistence, and mobility, atrazine is
found commonly in freshwater environments where many fish
and amphibians develop [10,11]. Atrazine has a variety of effects
on freshwater organisms, including fish and amphibians. For
example, atrazine has been reported to affect behavior [12,13],
physiology, and growth [14,15]; elevate mortality [16–18];
increase infections and suppresses immunity [19–22]; and induce
community-wide, indirect effects [23–26]. A recent meta-
analysis revealed that all of these effects appear to be general,
with the exception of elevated mortality [9].
More specifically, atrazine has been reported as both an
intraorganismal and an interorganismal infodisruptor. Atrazine
has been documented as an endocrine disruptor of freshwater
fish and amphibians [9,27,28]. In addition, several studies
provide evidence that exposure to ecologically relevant
concentrations of atrazine can reduce the olfactory abilities of
fish and crayfish [29–35]. For example, atrazine impaired
olfaction in salmon, with adverse effects on imprinting, homing,
and migratory behaviors [34]. Other studies have shown that
atrazine exposure decreased the response of male salmon to
reproductive priming hormones released in the urine of female
salmon [29,33] and decreased the response of goldfish to
chemical cues signaling predation [32]. Several fish studies
using electro-olfactograms, which measure the changing
electrical potentials of the olfactory epithelium, found that
short term, low-level exposure to atrazine reduced electro-
olfactogram responses and olfactory-mediated behav-
iors [30,31]. In contrast, olfaction of American toad (Anaxyrus
americanus) tadpoles was not significantly affected by
atrazine [36]. Additional studies are needed on amphibians to
assess whether atrazine generally has adverse effects on
olfaction in freshwater vertebrates. In the present study, we
evaluated whether exposing Cuban tree frog (Osteopilus
septentrionalis) tadpoles to the estimated environmental
concentration of atrazine affects their abilities to detect
chemical cues from predatory dragonfly larvae (Libellulidae
sp.) and to exhibit appropriate antipredator behaviors. Given
* Address correspondence to jasonrohr@gmail.com
Published online 22 January 2016 in Wiley Online Library
(wileyonlinelibrary.com).
DOI: 10.1002/etc.3377
Environmental Toxicology and Chemistry, Vol. 9999, No. 9999, pp. 1–6, 2016
#2016 SETAC
Printed in the USA
1
that there are more studies suggesting that atrazine is an
infodisruptor than not, we hypothesized that exposure to
atrazine would decrease tadpole detection of chemical cues
from predators.
Background on field concentrations of atrazine
Although atrazine has been banned in Europe, it remains one
of the most commonly used pesticides in the United States and
the rest of the world, where it is used heavily in corn and
sorghum production [9]. Atrazine is relatively persistent, with
an aqueous photolysis half-life of 335 d under natural light and
neutral pH and half-lives that can exceed 3 mo in mescosms [9].
Atrazine is also relatively mobile, regularly entering water
bodies with runoff and rain inputs [9]. Atrazine has been
detected in rain or air from European and US sites more than any
other currently used pesticide [37].
In the present study, we tested the effects of the estimated
environmental concentration (EEC) of atrazine on tadpole
olfaction. The EEC is the concentration estimated to enter a
standardized farm pond that is a standardized distance from the
application site (based on particular soil properties) [23]. The
EEC was calculated using US Environmental Protection
Agency (USEPA) GENEEC software (based on applications
to corn) and is the concentration used in registering
pesticides [23]. The measured concentration to which tadpoles
were exposed was 178 mg/L of atrazine.
To place this concentration in an ecological context, we
provide background on measured concentrations of atrazine in
the environment. The maximum reported wet deposition of
atrazine is 154 mg/L from Iowa precipitation [38]. Data on
atrazine concentrations in surface water are more abundant for
lotic (streams and rivers) than lentic (lakes, ponds, wetlands,
ditches) systems, primarily because of the extensive stream
monitoring conducted by the US Geological Survey National
Water Quality Assessment project [39]. Atrazine has been
detected in streams at 200 mg/L and repeatedly has been
detected in streams above 100 mg/L [39]. However, more
amphibians develop in lentic than lotic systems, where water is
not replenished and chemicals can concentrate as lentic systems
dry. Rohr and McCoy [9] reported that maximum reported
atrazine concentrations in lentic systems are often 2.5 to 10
times higher than maximum concentrations in lotic systems. For
example, atrazine concentrations in lentic surface waters have
been reported at 131 mg/L in Mississippi (USA) [40], 681 mg/L
in Ontario (Canada) [41], 850 mg/L in Florida (USA) [42],
1068 mg/L in Kansas (USA) [43], 1096 mg/L in Nebraska
(USA) [44], and 2300 mg/L in Iowa (USA) [45]. These findings
seem to belie probabilistic aquatic ecological risk assessments
for atrazine that suggest that concentrations near or above the
EEC are extremely rare [46] and demonstrate that measured
concentrations of atrazine near and well above the EEC are
widespread. In summary, the EEC is the concentration the
USEPA estimates to enter farm ponds regularly and is not a
worst-case scenario, because it was exceeded in at least 4 states
in the United States and in Canada. As such, the concentration of
atrazine we test in the present study is ecologically relevant
based both on regulatory standards and measured field
concentrations.
MATERIALS AND METHODS
Tadpole rearing
Cuban tree frog tadpoles were collected from the Botanical
Gardens at the University of South Florida in Tampa, Florida.
Once in the lab, tadpoles were maintained under laboratory
conditions at 22 8C and a 12:12-h light:dark cycle. Eighty
tadpoles (Gosner stages ranging from 25 to 41; mean standard
error [SE], 34.00 0.53) were kept individually in 250 mL of
pond water in a standard 0.47-L glass mason jar. One-half the
tadpoles were then exposed to 7.81 mL of either acetone or a
3.26-mg/mL stock solution of atrazine dissolved in acetone, to
achieve an estimated actual concentration of 178 mg/L of
atrazine (measured with an Abraxis enzyme-linked immuno-
sorbent assay microtiter plate kit; Abraxis). This is the estimated
environmental concentration of atrazine for farm ponds.
Tadpoles were exposed to acetone control or atrazine for 7 d.
They were fed agarose discs containing spirulina and flaked fish
food ad libitum, and we checked survival daily.
Predator cue collection
Aquatic dragonfly larvae regularly prey on frog tadpoles [24]
and are often the dominant predator in temporary pond
ecosystems. We collected 12 Libellulidae dragonfly larvae
from Spectrum Pond at the University of South Florida and
maintained them in an 11.36-L aquarium filled with 5.68 L
of dechlorinated tap water for 24 h. This tank also contained
12 Cuban tree frog tadpoles to provide food for the dragonfly
larvae. The water in this tank served as our predator cue source.
Behavioral observations
For behavioral observations, 20 plastic shoeboxes were laid
out in a 5-by-4 grid, and each was filled with 2 L of pond water.
Observations were double blind, as the observer did not have
access to the treatment groups of the individual tadpoles at the
time of the observations. One tadpole was placed in each
shoebox; 10 of the tadpoles were exposed to acetone control,
and the other 10 were exposed to atrazine. Scan samples were
conducted for 8 min before cues were added, recording the
location of tadpoles in each shoebox (left or right side of
the arena) and whether individual tadpoles were moving.
After the initial 8-min period, a repeat pipetter was used to
add 3 mL of either a control (water) or experimental predator
cue to 1 side of the container, while an equal amount of water
was added to the opposite side of the container. Cue and control
solutions were added carefully and simultaneously so that
the liquid ran down the side of the container to minimize
disturbance. One-half the predator cue addition in each
treatment occurred on the right side of the container, and
one-half was added to the left side. Thus, one-half the atrazine-
exposed and one-half the acetone-exposed tadpoles received
predator cue on 1 side of the container and the other tadpoles
received water on both sides of the container. Having water on
both sides allowed us to quantify activity levels in the absence of
predator cue.
Immediately after cue additions, which took approximately
3 min, we performed scan samples for 20 min, again recording
the location and activity of tadpoles. Given that 1 experimenter
could only observe 20 tadpoles at a time, and we wanted the
same experimenter making all observations, we conducted 3
additional temporal blocks in sequence, so that all 80 tadpoles
were exposed to the cues in a timeframe of approximately 4 h.
Immediately after observations in the shoeboxes were complete,
all tadpoles were returned to their mason jars containing their
assigned chemical treatment. The next day, the same procedure
was repeated for all 80 tadpoles, except that any tadpole that was
exposed to the predator cue on day 1 received water on both
sides of the arena on day 2 and vice versa. This experimental
design allowed us to use each tadpole as its own internal control
2Environ Toxicol Chem 9999, 2016 M. Ehrsam et al.
for both activity and locational preference. After the 2 d of
observations were complete, tadpoles were euthanized with
MS-222 solution. We then recorded the snout–vent length, mass
(g), and Gosner developmental stage [21] of each tadpole and
preserved tadpoles individually in 70% ethanol.
To determine the mean time it took the predator chemical
cues to diffuse across the test chamber, in separate trials, we
added 5 drops of food coloring to a randomly selected end of
5 test chambers each containing a tadpole. Time was recorded
when the dye had diffused to the halfway point of the tank and
again when the dye reached the opposite end of the tank.
Data analysis
We first transformed the data, giving observations on the
same side of the predator cue a value of 1 and on the opposite
side as the cue a value of –1. Hence, negative values refer to
avoidance and positive values refer to attraction. If a tadpole
was moving during an observation, it received a 1, and if it was
not moving it received a 0. Hence, the average of the activity
data reflects the proportion of observations where the tadpole
was moving.
Next, we conducted separate 2-way analyses of variance
(ANOVAs) on average location and activity before any
olfactory cues were added, testing for main and interactive
effects of the atrazine and predator treatments. This allowed us
to determine whether there were any differences between
treatments before the predator cues were added. We knew that
the predator cue would eventually diffuse across the entire test
arena within the 30-min trial. Thus, we were expecting to
detect an atrazine-by-predator-by-time interaction, because we
hypothesized that the solvent-exposed but not atrazine-exposed
animals would avoid the predator chemical cue early in the trial
when there was a clear gradient, but avoidance would eventually
subside once the cue fully dispersed. To conduct these analyses,
we averaged the location and activity data for each of the
four 5-min intervals of the 20-min trials. This enhanced our
likelihood of meeting the assumptions of ANOVA (because of
the central limit theorem). We then conducted a repeated
measures ANOVA with individual as a random effect (so we
could compare location and activity of each individual in the
presence and absence of predator cues) and tested for the main
and interactive effects of atrazine treatment, predator treatment,
and time on location and activity responses. Thirteen tadpoles
died during this experiment and thus were excluded from any
statistical analyses. All analyses were conducted using Statistica
6.0 (Statsoft).
RESULTS
Before any predator cues were added, the tadpoles showed
no significant difference in their average location in the atrazine
or predator cue treatments (p>0.05; Table 1 and Figure 1A).
After the predator cue was introduced, tadpoles exposed to
solvent, but not those exposed to atrazine, significantly avoided
the predator chemical cue for the first 10 min of observations but
exhibited no avoidance during the last 10 min (Figure 2). This
Table 1. Results of analyses of variance on tadpolelocation before and after
predator cue additions
Effect df F p
Pre-predator cue
Intercept 1 1.64 0.204
Atrazine 1 0.74 0.392
Error 67
Predator 1 0.02 0.883
Predator atrazine 1 0.44 0.507
Error 67
Post-predator cue
Intercept 1 0.21 0.648
Atrazine 1 1.82 0.182
Error 67
Time 3 0.37 0.778
Time atrazine 3 0.76 0.519
Error 201
Predator 1 0.67 0.416
Predator atrazine 1 0.12 0.730
Error 67
Time predator 3 2.10 0.102
Time predator atrazine 3 3.20 0.024
Error 201
Figure 1. Effect of atrazine on mean (1 standard error [SE]) tadpole
location before predator cue addition (A). Effect of atrazine (B) and predator
cue (C) treatments on mean (1 SE) activity of tadpoles before and after
predator cue introductions.
Atrazine reduces detection of predator chemical cues Environ Toxicol Chem 9999, 2016 3
resulted in a significant atrazine-by-predator cue-by-time
interaction (F
3,201
¼3.20, p¼0.024; Table 1). It took an
average of 8.8 0.97 min (SE; n¼5) for the food coloring to
diffuse fully across the test chamber. Hence, tadpole avoidance
of the predator cue seemed to last for approximately as long as
the cue gradient persisted.
Before any predator cues were added, tadpoles exposed to
atrazine were significantly more active than tadpoles exposed to
solvent control (F
1,67
¼7.84, p¼0.007; Figure 1B); but activity
levels did not differ significantly between tadpoles assigned to
receive predator cue or not (Table 2). After the predator cue
was introduced, tadpoles exposed to atrazine remained signi-
ficantly more active than tadpoles exposed to solvent control
(F
1,67
¼7.13, p¼0.009; Figure 1B). The tadpoles, however, did
not significantly reduce their activity in response to the predator
cue (Figure 1C) but did tend to exhibit less activity later than
earlier in the trial (F
3,201
¼4.45, p¼0.005; Figure 1C). There
were no significant statistical interactions on tadpole activity
(Table 2).
DISCUSSION
We found evidence that atrazine exposure impairs the ability
of Cuban tree frog tadpoles to detect chemical cues from
predators. Tadpoles exposed to solvent control significantly
avoided predator chemical cues, whereas tadpoles exposed
to atrazine did not (Figure 2). In addition, we revealed that
exposure to ecologically relevant concentrations of atrazine
caused hyperactivity in Cuban tree frog tadpoles compared with
tadpoles exposed to solvent control (Figure 1B). These results
have many similarities with other studies examining the effects
of atrazine on the behavior and olfactory abilities of amphibians
and fish.
Several other studies have shown that ecologically relevant
concentrations of atrazine affect the motor activity of fish and
amphibians [9]. After just 1 d of exposure to environmentally
relevant concentrations of atrazine, red drum fish larva
exhibited hyperactivity in the form of high swimming speeds,
erratic swimming paths, and increased energy use and metabolic
rates [47]. Another study investigating the sublethal effects of
atrazine in goldfish found that short-term exposures to low
atrazine concentrations caused activity increases, including
elevated burst swimming and surfacing activities [32]. Sala-
mander larva exposed to various concentrations of atrazine
exhibited higher activity than control larvae [11,14], and these
effects persisted for several months after atrazine exposure
ceased [12,13].
Our finding that atrazine exposure impairs the ability of
Cuban tree frog tadpoles to detect chemical cues that predators
release is consistent with several studies showing that ecolo-
gically relevant concentrations of atrazine can compromise the
olfactory abilities of fish and crayfish [29–35]. However, the
present study’s results are inconsistent with the only other study
that examined atrazine-related infodisruption in tadpoles [36].
Importantly, every study that detected atrazine-associated
infodisruption tested for infodisruption while the fish, crayfish,
or amphibian was being exposed to atrazine [29–35]. The single
study that did not detect infodisruption associated with atrazine-
exposed tadpoles transferred the tadpoles to atrazine-free water,
provided a 30-min acclimation period, and then introduced
chemical cues to elicit behavioral responses [36]. If the effect of
atrazine on the olfactory organs of vertebrates is short-lived,
Figure 2. The relationship between time since predator (Pred) cue addition and exposure to atrazine (Atr) or solvent (Solv) on the mean location (1 standard
error [SE] to reduce overlap of SE bars) of tadpoles. Positive values represent attraction to the predator cue, and negative values represent avoidance of the
predator cue.
Table 2. Results of analyses of variance on tadpole activity before and after
predator cue additions
Effect df F p
Pre-predator cue
Intercept 1 370.63 <0.001
Atrazine 1 7.84 0.007
Error 67
Predator 1 0.30 0.583
Predator atrazine 1 1.04 0.313
Error 67
Post-predator cue
Intercept 1 167.63 <0.001
Atrazine 1 7.13 0.009
Error 67
Time 3 4.45 0.005
Time atrazine 3 0.29 0.834
Error 201
Predator 1 2.05 0.157
Predator atrazine 1 2.67 0.107
Error 67
Time predator 3 0.51 0.674
Time predator atrazine 3 0.45 0.717
Error 201
4Environ Toxicol Chem 9999, 2016 M. Ehrsam et al.
then 30 min exposure to clean water might have been sufficient
time to rinse the atrazine from the olfactory organ, allow for at
least partial recovery of olfactory function, or prevent the
detection of any atrazine-induced olfactory impairment.
However, we cannot rule out species-level differences in
sensitivity to atrazine or other differences among the experi-
ments as explanations for the differences in their results. We
encourage future studies to evaluate species-level variation in
olfactory sensitivity to atrazine and how both durations and
concentrations of atrazine exposure affect amphibian olfaction.
Amphibians are in decline globally [48,49], and although it is
theoretically possible that atrazine-induced infodisruption could
contribute to these declines, we caution against extrapolating
these results to the population viability of amphibians in the wild
for several reasons. First, if the effects of atrazine are found to be
short-lived, it might suggest that any infodisruption might not
have substantial impacts on wildlife populations. Hence,
quantifying the duration of atrazine effects on olfaction would
offer insight into both discrepancies in the results among studies
and the likelihood that any infodisruption would have
population-level impacts. Second, if the olfaction of predators
is equally or more impaired by atrazine than the olfaction of
amphibian prey, then any infodisruption could benefit the prey
more than the predators. It is important, therefore, to understand
the net effects of pesticides on species’interactions before
reliably extrapolating to effects in nature [19]. Finally, there are
many factors that are not captured in a laboratory study that
could affect the effects of atrazine in the wild, such as variation
in temperature and organic material on which pesticides can
bind [18]. Regardless of how any atrazine-induced infodisrup-
tion influences populations of vertebrates in the wild, the present
study’s results contribute to the weight of evidence that atrazine
is, indeed, an interorganismal infodisruptor of vertebrates.
We encourage further studies to more thoroughly quantify
the consequences of such infodisruption and whether they are
capable of contributing to declines of biodiversity.
Acknowledgment—We thank B. Roznik and C. Gabor for help and support.
This research was supported by grants from the National Science
Foundation (EF-1241889), National Institutes of Health (R01GM109499,
R01TW010286), US Department of Agriculture (NRI 2006-01370, 2009-
35102-0543), and US Environmental Protection Agency (CAREER
83518801) to J. Rohr.
Data availability—Data are available by contacting the corresponding
author (jasonrohr@gmail.com).
REFERENCES
1. Lurling M, Scheffer M. 2007. Info-disruption: Pollution and the transfer
of chemical information between organisms. Trends Ecol Evol
22:374–379.
2. Schmidt KA, Dall SRX, van Gils JA. 2010. The ecology of information:
An overview on the ecological significance of making informed
decisions. Oikos 119:304–316.
3. Kidd KA, Blanchfield PJ, Mills KH, Palace VP, Evans RE, Lazorchak
JM, Flick RW. 2007. Collapse of a fish population after exposure to a
synthetic estrogen. P Natl Acad Sci USA 104:8897–8901.
4. Rohr JR, Madison DM, Sullivan AM. 2002. Sex differences and
seasonal trade-offs in response to injured and non-injured conspecifics
in red-spotted newts, Notophthalmus viridescens.Behav Ecol Sociobiol
52:385–393.
5. Chivers DP, Smith RJF. 1998. Chemical alarm signalling in aquatic
predator-prey systems: A review and prospectus. Ecoscience
5:338–352.
6. Rohr JR, Park D, Sullivan AM, McKenna M, Propper CR, Madison
DM. 2005. Operational sex ratio in newts: Field responses and
characterization of a constituent chemical cue. Behav Ecol 16:286–
293.
7. Rohr JR, Madison DM, Sullivan AM. 2003. On temporal variation and
conflicting selection pressures: A test of theory using newts. Ecology
84:1816–1826.
8. Kats LB, Dill LM. 1998. The scent of death: Chemosensory assessment
of predation risk by prey animals. Ecoscience 5:361–394.
9. Rohr JR, McCoy KA. 2010. A qualitative meta-analysis reveals
consistent effects of atrazine on freshwater fish and amphibians.
Environ Health Persp 18:20–32.
10. Knutson MG, Richardson WB, Reineke DM, Gray BR, Parmelee JR,
Weick SE. 2004. Agricultural ponds support amphibian populations.
Ecol Appl 14:669–684.
11. Rohr JR, Elskus AA, Shepherd BS, Crowley PH, McCarthy TM,
Niedzwiecki JH, Sager T, Sih A, Palmer BD. 2003. Lethal and sublethal
effects of atrazine, carbaryl, endosulfan, and octylphenol on the
streamside salamander, Ambystoma barbouri.Environ Toxicol Chem
22:2385–2392.
12. Rohr JR, Palmer BD. 2005. Aquatic herbicide exposure increases
salamander desiccation risk eight months later in a terrestrial
environment. Environ Toxicol Chem 24:1253–1258.
13. Rohr JR, Palmer BD. 2013. Climate change, multiple stressors, and the
decline of ectotherms. Conserv Biol 27:741–751.
14. Rohr JR, Elskus AA, Shepherd BS, Crowley PH, McCarthy TM,
Niedzwiecki JH, Sager T, Sih A, Palmer BD. 2004. Multiple stressors
and salamanders: Effects of an herbicide, food limitation, and
hydroperiod. Ecol Appl 14:1028–1040.
15. Larson DL, McDonald S, Fivizzani AJ, Newton WE, Hamilton SJ.
1998. Effects of the herbicide atrazine on Ambystoma tigrinum
metamorphosis: Duration, larval growth, and hormonal response. Phys
Zool 71:671–679.
16. Rohr JR, Sager T, Sesterhenn TM, Palmer BD. 2006. Exposure,
postexposure, and density-mediated effects of atrazine on amphibians:
Breaking down net effects into their parts. Environ Health Persp
114:46–50.
17. Storrs SI, Kiesecker JM. 2004. Survivorship patterns of larval
amphibians exposed to low concentrations of atrazine. Environ Health
Persp 112:1054–1057.
18. Rohr JR, Sesterhenn TM, Stieha C. 2011. Will climate change
reduce the effects of a pesticide on amphibians? Partitioning effects
on exposure and susceptibility to pollution. Glob Change Biol
17:657–666.
19. Rohr JR, Raffel TR, Sessions SK, Hudson PJ. 2008. Understanding the
net effects of pesticides on amphibian trematode infections. Ecol Appl
18:1743–1753.
20. Kerby JL, Storfer A. 2009. Combined effects of atrazine and
chlorpyrifos on susceptibility of the tiger salamander to Ambystoma
tigrinum virus. EcoHealth 6:91–98.
21. Rohr JR, Schotthoefer AM, Raffel TR, Carrick HJ, Halstead N,
Hoverman JT, Johnson CM, Johnson LB, Lieske C, Piwoni MD, Schoff
PK, Beasley VR. 2008. Agrochemicals increase trematode infections in
a declining amphibian species. Nature 455:1235–1239.
22. Rohr JR, Raffel TR, Halstead NT, McMahon TA, Johnson SA,
Boughton RK, Martin LB. 2013. Early-life exposure to a herbicide has
enduring effects on pathogen-induced mortality. P Roy Soc Lond B Bio
280:20131502.
23. Halstead NT, McMahon TA, Johnson SA, Raffel TR, Romansic JM,
Crumrine PW, Rohr JR. 2014. Community ecology theory predicts the
effects of agrochemical mixtures on aquatic biodiversity and ecosystem
properties. Ecol Lett 17:932–941.
24. Rohr JR, Crumrine PW. 2005. Effects of an herbicide and an insecticide
on pond community structure and processes. Ecol Appl 15:1135–1147.
25. Staley ZR, Rohr JR, Harwood VJ. 2011. Test of direct and indirect
effects of agrochemicals on the survival of fecal indicator bacteria. Appl
Environ Microbiol 77:8765–8774.
26. Staley ZR, Rohr JR, Senkbeil JK, Harwood VJ. 2014. Agrochemicals
indirectly increase survival of E. coli O157: H7 and indicator bacteria
by reducing ecosystem services. Ecol Appl 24:1945–1953.
27. Hayes TB, Khoury V, Narayan A, Nazir M, Park A, Brown T, Adame L,
Chan E, Buchholz D, Stueve T, Gallipeau S. 2010. Atrazine induces
complete feminization and chemical castration in male African clawed
frogs (Xenopus laevis). P Natl Acad Sci USA 107:4612–4617.
28. Langlois VS, Carew AC, Pauli BD, Wade MG, Cooke GM, Trudeau
VL. 2010. Low levels of the herbicide atrazine alter sex ratios and
reduce metamorphic success in Rana pipiens tadpoles raised in outdoor
mesocosms. Environ Health Persp 118:552–557.
29. Moore A, Waring CP. 1998. Mechanistic effects of a triazine pesticide
on reproductive endocrine function in mature male Atlantic salmon
(Salmo salar L.) parr. Pestic Biochem Phys 62:41–50.
Atrazine reduces detection of predator chemical cues Environ Toxicol Chem 9999, 2016 5
30. Tierney KB, Ross PS, Jarrard HE, Delaney KR, Kennedy CJ. 2006.
Changes in juvenile coho salmon electro-olfactogram during and after
short-term exposure to current-use pesticides. Environ Toxicol Chem
25:2809–2817.
31. Tierney KB, Singh CR, Ross PS, Kennedy CJ. 2007. Relating olfactory
neurotoxicity to altered olfactory-mediated behaviors in rainbow
trout exposed to three currently-used pesticides. Aquat Toxicol
81:55–64.
32. Saglio P, Trijasse S. 1998. Behavioral responses to atrazine and diuron
in goldfish. Arch Environ Contam Toxicol 35:484–491.
33. Moore A, Lower N. 2001. The impact of two pesticides on olfactory-
mediated endocrine function in mature male Atlantic salmon (Salmo
salar L.) parr. Comp Biochem Phys B 129:269–276.
34. Moore A, Lower N, Mayer I, Greenwood L. 2007. The impact
of a pesticide on migratory activity and olfactory function in
Atlantic salmon (Salmo salar L.) smolts. Aquaculture 273:350–
359.
35. Belanger RM, Peters TJ, Sabhapathy GS, Khan S, Katta J, Abraham
NK. 2015. Atrazine exposure affects the ability of crayfish (Orconectes
rusticus) to localize a food odor source. Arch Environ Contam Toxicol
68:636–645.
36. Rohr JR, Swan A, Raffel TR, Hudson PJ. 2009. Parasites, info-
disruption, and the ecology of fear. Oecologia 159:447–454.
37. van Dijk HFG, Guicherit R. 1999. Atmospheric dispersion of current-
use pesticides: A review of the evidence from monitoring studies. Water
Air Soil Poll 115:21–70.
38. Hatfield JL, Wesley CK, Prueger JH, Pfeiffer RL. 1996. Herbicide and
nitrate distribution in central Iowa rainfall. J Environ Qual 25:259–
264.
39. US Geological Survey. National Water Quality Assessment (NAWQA)
Program. cited 2015 November 24]. Available from: http://cida.usgs.
gov/nawqa_www/nawqa_data_redirect.html?p¼nawqa
40. Kroger R, Moore MT, Brandt JR. 2012. Current- and past-use pesticide
prevalence in drainage ditches in the Lower Mississippi Alluvial
Valley. Pest Manag Sci 68:303–312.
41. Frank R, Braun HE, Ripley BD, Clegg BS. 1990. Contamination of
rural ponds with pesticide, 1971–85, Ontario, Canada. Bull Environ
Contam Toxicol 44:401–409.
42. Graves GA, Wan YS, Fike DL. 2004. Water quality characteristics of
storm water from major land uses in South Florida. J Am Water Resour
As 40:1405–1419.
43. Kadoum AM, Mock DE. 1978. Herbicide and insecticide residues in
tailwater pits: Water and pit bottom soil from irrigated corn and
sorghum fields. J Agri Food Chem 26:45–50.
44. Watanabe H, Watermeier NL, Steichen JM, Barnes P, Phong TK. 2007.
Impacts of tillage and application methods on atrazine and alachlor
losses from upland fields. Weed Biol Manag 7:44–54.
45. Kolpin DW, SneckFahrer D, Hallberg GR, Libra RD. 1997. Temporal
trends of selected agricultural chemicals in Iowa’s groundwater, 1982–
1995: Are things getting better? J Environ Qual 26:1007–1017.
46. Giddings JM, Anderson TA, Hall LW, Hosmer AJ, Kendall RJ,
Richards RP, Solomon KR, Williams WM. 2005. Atrazine in North
American Surface Waters: A Probabilistic Aquatic Ecological Risk
Assessment. Society for Envrionmental Toxicology and Chemistry,
Pensacola, FL, USA.
47. Alvarez MD, Fuiman LA. 2005. Environmental levels of atrazine and
its degradation products impair survival skills and growth of red drum
larvae. Aquat Toxicol 74:229–241.
48. Rohr JR, Raffel TR. 2010. Linking global climate and temperature
variability to widespread amphibian declines putatively caused by
disease. P Natl Acad Sci USA 107:8269–8274.
49. Wake DB, Vredenburg VT. 2008. Are we in the midst of the sixth mass
extinction? A view from the world of amphibians. P Natl Acad Sci USA
105:11466–11473.
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