EFFECTS OF ATRAZINE EXPOSURE
ON AROMATASE EXPRESSION IN
MALE ZEBRAFISH (DANIO RERIO)
Carrie Schmaus Senior Honors Thesis MAY 16, 2015 DEPARTMENT OF BIOLOGY Wittenberg University, Springfield, Ohio Schmaus 1
“Science is a principle and a process of seeking truth. Truth cannot be purchased, and, thus,
truth cannot be altered by money.” –Tyrone Hayes
Abstract
Atrazine, the second most utilized herbicide in the United States, has been shown to
disrupt endocrine systems in a variety of organisms. Studies have shown atrazine to cause male
frogs to become hermaphrodites, as well as inflict chromosomal damage to the ovary cells of
hamsters and induce false pregnancies in rats. However, other studies have refuted these results,
showing that legal limits of atrazine have negligible effects on exposed organisms. The topic of
atrazine use has recently become controversial, especially as the product benefits the U.S.
agricultural market by billions of dollars every year. Most studies performed on the effects of
atrazine exposure have been conducted on frogs, so these results are well documented. However,
zebrafish, a vertebrate model organism, has been less studied. By exposing male zebrafish to an
environmentally relevant concentration of atrazine, this study aimed to elucidate the effects of
atrazine on the expression of aromatase. Aromatase is the enzyme responsible for both sex
determination and conversion of testosterone to estrogen in zebrafish, though the mechanism by
which it is synthesized is poorly understood. Results showed that atrazine exposed organisms did
not exhibit aromatase expression, though non-exposed organisms did. These results may be due
to experimental errors, issues with the experimental design (in that the atrazine concentration
used and/or the exposure period were not sufficient), or the possibility that atrazine does not
influence the expression of aromatase, but instead, affects the activity of the enzyme. Further
research is recommended to elucidate the effects of environmentally relevant concentrations of
atrazine on male zebrafish.
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Introduction
Atrazine
Atrazine, a commercial triazine herbicide (2-chloro-4-ethylamino-6-isopropyl-amino-striazine), is mainly used to control broadleaf weeds and grasses, most often in corn crops
(Solomon et al., 1995). Atrazine is usually applied directly to the leaves of the intended target
weeds in large scale agricultural practices, though it is recommended that it be applied before
corn reaches a foot in height; this application technique classifies atrazine as a post-emergence
herbicide (Hager and McGlamery, 1997; Home Depot, 2015). This recommendation implies that
corn will be not be considered a broadleaf plant until after it has grown over a foot tall. Atrazine
had been used all over the world, and is applied heavily in the United States currently; about 80
million pounds per year (Hayes et al., 2010). The Midwest accounts for much of this atrazine
application due to the importance of agriculture in this region of the country. Illinois alone
accounts for 17% of all use in the United States, followed by Iowa (14%), Nebraska (11%), and
Indiana (10%) (Solomon et al., 1995). Atrazine is currently manufactured by the company
Syngenta, and contributes to the economic success of American farmers. Because of its
commercial value, Atrazine has become a topic of not only ecological discussion, but a political
and economic topic as well.
The Atrazine Controversy
Atrazine was first patented in Switzerland in 1958, and was patented for use in the United
States in 1959 (Solomon et al., 1995). Since then, the use of atrazine has increased, until studies
were conducted that showed atrazine to be detrimental to the sexual health of exposed
organisms. Finally, in 1994, the EPA began to investigate the herbicide, and studies were
conducted to determine the safety of atrazine (Solomon et al., 1995). Though there have been
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many reports showing detrimental effects of the herbicide, conflicting studies have also shown
effects on exposed organisms to be negligible at small concentrations. As such, the U.S.
continues to support the use of the herbicide. However, the European Union and Switzerland, the
country responsible for the herbicide, have both banned atrazine since such studies have surfaced
(NDRC, 2009).
Unlike Europe, the dispute concerning atrazine usage continues in the United States.
Tyrone Hayes, the man thought to have started the atrazine debate, is a professor at UC Berkley,
but recently has become a political figure in his public criticism of atrazine. As a scientist once
funded by Syngenta, Hayes found the product to be detrimental to the sexual development in
frogs (Hayes et al., 2003; Hayes et al., 2006 Hayes et al., 2010). As his research was published,
Hayes discontinued his employment with Syngenta and was given reason to believe the
corporation was aiming to discredit his work. These suspicions were confirmed with the courtdictated release of Syngenta’s internal documents, some of which were memos detailing ways to
involve Hayes in a scandal to discredit his research (Aviv, 2014).
Hayes continues to study atrazine effects in frogs and publish his findings, which have
both been confirmed by other independent scientists and denied by studies funded by Syngenta
and the EPA. Syngenta’s interest in keeping their products on the shelves is warranted: atrazine
supports up to 85,000 jobs and benefits farmers by $3 billion annually, as well as increasing crop
yields by 600 million bushels per year (Syngenta, 2012). The continued usage of atrazine is
important for the corporation and farmers alike, so research showing atrazine is not harmful to
the organisms that experience exposure protects the product. However, there is much reason to
believe that inherent bias and money involved in these research endeavors is clouding the truth.
The focus of this paper will not be the politics of atrazine use in the United States, but it is
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important to note that more factors are present in the discussion of atrazine than is typical for a
scientific endeavor.
Atrazine in Ground and Surface Water
Atrazine pollution of groundwater occurs when the herbicide leaches through the soil to
the aquifer, and is about 20 times more frequently found in groundwater than any other herbicide
(as cited in Graymore et al., 2001); a 2009 U.S. geological survey found atrazine to be present at
detectable levels in 40% of the groundwater sampled across the nation (NDRC, 2009). Of the 40
watersheds tested, a fourth of these had concentrations of atrazine above 1 ppb, while other
watersheds were shown to have peak concentrations exceeding 100 ppb (NRDC, 2009). Atrazine
is found in highest concentrations after storm events during the spring and summer months, often
corresponding with many mating events for organisms living in the surrounding ponds and
streams (Graymore et al., 2001; NRDC, 2009).
Atrazine enters surface water, such as streams, rivers, and lakes, primarily through runoff
(as cited in Graymore et al., 2001). The geological survey mentioned above found atrazine to be
present in detectable concentrations in 75% of the streams that were sampled, and concentrations
as high as 1000 ppb have been found in streams directly adjacent to crops where atrazine was
applied (NRDC, 2009; cited in Graymore et al., 2001). These extremely high concentrations are
only seen if atrazine application and a storm event happen simultaneously, however, and it
should be noted that concentrations in these same types of areas have also been found to be as
low as 0.2 ppb. Fluctuating concentrations of atrazine in groundwater are due in part to the
leaching potential of the soil and the amount of water moving through said soil, while
fluctuations in surface water are often due to the timing of storm events (Huddleston, 1996;
Graymore et al., 2001).
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Restrictions on Atrazine in the United States
The EPA has designated a maximum contaminant level (MCL) for atrazine in drinking
water at 3 ppb, or 0.003 mg/L—concentrations under this level are thought to be safe for human
consumption (EPA, 2013). However, it should be noted that 3 ppb is a running average, so high
influxes of atrazine, such as application events, may be averaged out by lower concentrations
that occur after the application event (NDRC, 2009). Additionally, the EPA only dictates
concentrations in the context of safe human exposure; effects on organisms living in streams that
experience atrazine exposure have not been factored into determining the MCL. Regardless, the
EPA re-registered atrazine in 2006 as a safe chemical, and the heavy use of atrazine in the United
States continues currently (Syngenta, 2010).
There are about 50,000 community drinking water systems in the U.S., 40,000 of which
are serviced by supplies sourced from groundwater (for example, well water). The remaining
10,000 are fed by surface water—lakes, streams, and rivers—and these sources are the focus of
EPA monitoring, because, according to the EPA, atrazine concentrations tend to be higher in
surface water than ground water (EPA, 2013). Of the 50,000 community drinking water sources,
200 have been classified as “high risk” by the EPA, due to their proximity to agricultural areas.
In these areas, weekly monitoring during growing season is conducted to determine if the
atrazine level is above permissible levels; only eight of these systems have been shown to rise
significantly above 3 ppb consistently. When levels rise above the acceptable 3 ppb, site-specific
mitigation plans are put into action, and atrazine use is restricted in these areas. However, the
EPA has not put any restrictions in places where the water is not part of the drinking water
system, so, again, water sources adjacent to agricultural sites serving as habitats for animals have
not been considered from a federal restriction standpoint.
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Atrazine and Animals
The main research done on atrazine has been conducted by the aforementioned Tyrone
Hayes, and his focus organism has been frogs. His groundbreaking study showed concentrations
as low as 0.2 ppb atrazine to produce American leopard frogs with hermaphroditic qualities
(Hayes et al., 2003). This study also found 25 ppb to cause decreased testosterone levels in frogs
exposed to atrazine throughout their larval development to metamorphosis. The testosterone
levels were found to be so decreased, in fact, that males were found to have lower testosterone
levels than females (Hayes et al., 2003). Additionally, in collecting and analyzing frogs from
areas of atrazine exposure, this study showed frogs collected from the field to exhibit gonadal
dysgenesis and hermaphroditism, results that were confirmed by a lab exposure of atrazine which
created the same dysfunctions in male frogs (Hayes et al., 2003).
However, there have also been many studies conducted to refute these results. One such
study performed in the lab found that exposure to 0.01, 0.1, 1, 25 100 ppb atrazine from dpf (day
post fertilization) 8 until completion of metamorphosis, or dpf 83, showed no differences from
the exposed group to the control group (Kloas et al., 2008). That is, atrazine exposure did not
affect growth, larval development, or sexual differentiation in African clawed frogs. These
results directly refute what was found by Hayes, potentially due to the difference in exposure
period, as well as the species of frogs, but studies such as this have been convincing enough for
the EPA to re-register atrazine as an accepted herbicide. However, it is very important to note
that the Kloas study was funded by Syngenta Crop Protection, which, as mentioned previously,
is the company that manufactures atrazine.
Though frogs have been the subject of heavy atrazine investigation, studies in other
species have shown atrazine to be detrimental across the animal kingdom. Studies on rats have
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shown that short-term exposure to atrazine at 0-200 mg/kg per day causes preimplantation loss
(complications with pregnancy), and at 50-300 mg/kg per day for up to 21 days caused 7/9 rats to
experience a false pregnancy (Cooper et al., 2000). Other studies have shown hamsters to
experience chromosomal damage in their ovary cells, as well as damaged bone marrow in mice
(as cited in Graymore et al., 2001). Additionally, farm workers that use atrazine were found to
experience chromosomal abnormalities in lymphocyte cultures, and chlorinated herbicides, such
as atrazine, have been shown to trigger breast cancer development in humans (as cited in
Graymore et al., 2001). Clearly, effects such as these warrant some investigation, especially for
the millions of people living in agricultural areas where atrazine is in heavy use.
Atrazine and Fish
Studies in fish have shown exposed organisms to exhibit behavioral changes, especially
in swimming behavior (as cited in Graymore et al., 2001). Trout have also been seen to exhibit
reduced motility, a change in appearance, and balance changes after exposure to atrazine (as
cited in Graymore et al., 2001). These changes in fish are often behavioral, though some changes
to their endocrine systems have been seen as well. For example, one study tested effects of
atrazine exposure on zebrafish. Zebrafish are regarded to be a model organism, and are used for
experiments because the effects shown in zebrafish can suggest implications for other fish, as
well as vertebrates as well. This particular study exposed zebrafish to 2.2-20 ppb atrazine, and
this low concentration was found to increase the ratio of female to male fish as well as to alter
hormone networks and normal endocrine development. The reasoning proposed by Suzawa and
Ingraham suggest that atrazine affects the aromatase pathway indirectly, a hypothesis that has
been proposed by others as well (2008).
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The Aromatase Hypothesis
The current theory is that atrazine is an inducer for aromatase, an enzyme that catalyzes
the conversion of testosterone to estradiol, which is a type of estrogen (Figure 1; Hayes et al.,
2001; “Aromatase”). In this pathway, atrazine is hypothesized to increase the activity or
expression of aromatase; whether aromatase activity (how active the aromatase proteins are) or
the aromatase expression (the actual volume of aromatase) is the mechanism effected by atrazine
is unknown. Regardless of the mechanism, an increase in aromatase activity or expression is
hypothesized to convert more testosterone to estradiol, resulting in lower testosterone levels and
higher estrogen levels in atrazine-exposed organisms. In a phenomenon that has been coined
feminization, some genetic males exposed to atrazine become hermaphroditic, because they
develop both male and female reproductive parts (Hayes et al., 2006).
Figure 1. Conversion pathway of testosterone to estradiol, an estrogen. Note that aromatase
catalyzes this reaction (Wikipedia Commons).
Accordingly, other studies have shown that exposure to an antiandrogen (testosterone
inhibitor) or exogenous estrogen resulted in the same type of feminization effects in frogs as
atrazine exposure (Hayes et al., 2006). This phenomenon offers support for what is called the
“aromatase hypothesis”, which hypothesizes that atrazine-exposed organisms experience a
combination of increased estrogen and reduced testosterone (androgen) levels due to the
increased activity or expression of aromatase. Other studies cited in this same article have shown
that atrazine increases aromatase activity or expression in frogs, though they were unable to
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determine the exact mechanism due to inconsistent results. However, the combination of
depleting androgens and increasing estrogens has been shown to cause demasculinization and
feminization in many organisms, including amphibians, fish, reptiles, and mammals. Currently,
scientists believe the aromatase hypothesis to be a likely explanation (Hayes et al., 2006).
Atrazine and zcyp19a1
To understand the mechanism by which atrazine is hypothesized to increase the
expression or activity of aromatase, the mechanism of molecular expression must also be
understood. In eukaryotes, such as zebrafish, transcription occurs in the nucleus and produces,
among other products, mRNA. mRNA goes through a process called translation, in which it
serves as a template for a string of amino acids, or protein (Alberts et al, 1998). Regulation of
these processes often depends on transcription factors, as specific factors must be present for
transcription to take place properly. Genes must be activated by receptors to produce the protein,
or string of amino acids, that is made after transcription and translation. Thus, the receptor’s
activity is important in determining how much protein will be made, which affects the cell’s
function.
The exact molecular mechanisms that atrazine effects are generally unknown, but one
study has shown atrazine to cause an increase in expression of zcyp19a, the gene that encodes for
aromatase (Suzawa and Ingraham, 2008). In zebrafish, there are actually two genes that encode
for different types of aromatase: zcyp19a1 and zcyp19a2. zcyp19a1 encodes for gonadal
aromatase, and has been shown to be upregulated by atrazine (gene expression increases in the
presence of atrazine). The other gene, zycp19a2, is an estrogen-responsive gene that encodes for
aromatase in the brain, and atrazine was not shown to alter the expression of this gene (Suzawa
and Ingraham, 2008).
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Suzawa and Ingraham showed elevated expression of zycp19a1 with atrazine exposure,
but rejected the idea that atrazine directly causes this increased expression (2008). Instead, the
authors proposed an indirect method by which zycp19a1 expression was increased by atrazine
(ultimately increasing the activity or expression of aromatase). Here, atrazine is hypothesized to
stimulate the NR5A family of receptors, which then causes increased transcription of the
zycp19a1 gene. Because the zycp19a1 gene encodes for gonadal aromatase, an increase in
expression of this gene would cause elevated activity or expression of the aromatase protein
(Suzawa and Ingraham, 2008; Figure 2).
Figure 2. Proposed mechanism of
aromatase expression. Atrazine is
hypothesized to active the NR5A family
of receptors, therefore, indirectly
increasing the expression of aromatase
(Adapted from Suzawa and Ingraham
2008).
This study has highlighted one potential mechanism of how atrazine increases the expression or
activity of aromatase, but more research is needed to definitively understand the effects of
atrazine on this pathway. This study aimed to begin to elucidate some of the effects of atrazine
on the aromatase pathway in male zebrafish.
Experimental Design
To continue to determine atrazine’s effects on freshwater organisms, male zebrafish were
exposed to 70 pbb atrazine for nine days. 70 ppb was chosen because even though this
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concentration is much above the EPA’s drinking water standards, it has been found in water
bodies adjacent to fields after atrazine is applied and there is a storm (EPA, 2013; NRDC, 2009).
Zebrafish were utilized in this study because they serve as a model vertebrate organism, and
research on the effects of atrazine on the aromatase pathway in male zebrafish have only
undergone preliminary investigation. The nine day exposure period was decided upon because
zebrafish complete spermatogenesis in 6 days, so a nine-day exposure period would ensure that
at least one cycle of spermatogenesis was exposed to atrazine (Leal et al., 2009). Because
atrazine is hypothesized to demasculinize male organisms, sperm counts were a parameter of
investigation during this study (Hayes et al., 2010). The mechanism of aromatase expression or
activity increase was also investigated, though, due to budget constraints, only aromatase
expression was analyzed in this study. Here, the aromatase hypothesis was accepted as an
underlying assumption, and the hypothesis was that atrazine exposure would increase aromatase
expression in the exposed group of zebrafish when compared to the non-exposed group.
Materials and Methods
Exposure
Adult zebrafish (Danio rerio) were split into two groups: a non-exposed group (5 male, 2
female fish in fish water) and an exposed group (4 male, 2 female fish exposed to 70 ppb
commercially available Atrazine4 (Sunniland)) (3.5 µL Atrazine4 + 2 liters fish water). One liter
was emptied each day and replaced with one liter of fish water for the non-exposed group and
one liter of 70 ppb atrazine for the exposed group (following Kazeto et al., 2004). However, due
to a math error in which 1.25 µL of atrazine was added to the replacement liter in the exposed
group rather than the correct 1.75 µL, both tanks were completely emptied during the third day
and refilled with the correct concentrations.
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Experimental conditions were kept as consistent as possible, with both tanks being
exposed to identical humidity and light/dark schedule (14 hours light, 10 hours dark) and a
temperature of 27-280C. Each tank was fed twice per day (in the morning between 10 am – 1 pm
and in the evening between 7:30-8:45), except for the last day of exposure, when both tanks were
only fed in the evening. The exposure period lasted 9 days (Table 1).
Date
02.17.15
02.18.15
02.19.15*
02.20.15
02.21.15
02.22.15
02.23.15
02.24.15
02.25.15
1st Feeding Time
11:15 am
10:10 am
11:30 am
12:50 pm
11:30 am
11:30 am
12:30 pm
11:20 am
---
2nd Feeding Time
8:00 pm
8:30 pm
7:30 pm
7:45 pm
8:45 pm
8:15 pm
8:30 pm
8:45 pm
8:45 pm
Temperature (0C)
27
27
27
27
27
27
27
28
27
Table 1. Dates, feeding times, and temperature of room for exposure period.
*Water was fully changed on this day.
At the end of the exposure period, fish were anesthetized in 2% Tricane (Sigma Aldrich,
E10521; 2% Tricane + fish water) for sperm isolation, and eventually sacrificed by a lethal dose
of anesthetic to conduct the remainder of the experiment.
Sperm Isolation
Sperm isolation was attempted on all non-exposed males once before the official
experiment began, and again after the experiment had commenced. Exposed males only
experienced one attempted sperm isolation. To isolate sperm, fish were anesthetized in 2%
Tricane solution. Fish were patted dry with a paper towel (as water activates sperm), and placed
in a damp sponge containing a slit big enough for a fish (Figure 3).
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Figure 3. Schematic of sperm
isolation, zebrafish represented in
green. Note anal fins must be
separated to access anal pore.
The fish was placed upside down and the anal fins were separated. Under a dissecting
microscope (Leica Zoom 2000), a 0.2 µL microcapillary tube (Sigma Aldrich, P1549) was
placed in the anal pore while the fish was massaged with forceps to extract the sperm (following
Morris et al., 2003).
Isolated sperm was placed on a hemocytometer (Digital Bio, DHC-N01) slide. During the
initial trial, there was not enough sperm to reach the grid on the slide. By adding 1 µL of Hank’s
final solution to the extracted sperm, the sperm could reach the grid and the sperm could be counted
(Hank’s final solution; Westerfield, 2000).
Testes Isolation
Testes were isolated following a protocol for zebrafish organ dissection (Gupta and
Mullins, 2010). In short, euthanized fish were patted dry, placed under a dissection microscope,
and an incision was made anterior to the anal fin through the belly up to the operculum. The
operculum and pectoral fin were removed to expose the gills, and an incision posterior the gills
down to the anal fin through the belly was made. The skin and underlying muscle was removed,
and the testes were removed using forceps. The testes appear as long, white paired organs
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attached to the dorsal body wall. These were placed in homogenizers and kept on dry ice to
reduce the amount of RNA degradation. A total of two groups of testes were isolated (nonexposed and exposed). Testes belonging to the same group were homogenized together to
achieve the maximum amount of RNA.
RNA Isolation
RNA was isolated following the TRIzol protocol (Life Technologies, 15596-026). In
short, testes were homogenized with a tissue-grinding mortar and pestle in a fume hood. 1 mL of
TRIzol was added for each group of testes, homogenized, transferred to a microcentrifuge tube,
and incubated at room temperature for 5 minutes. 0.2 mL of chloroform was added and mixed
via vigorous inversion about 15 times, and incubated at room temperature for 2-3 minutes.
Homogenate was centrifuged at 12,000 x g for 15 minutes at 40C. The aqueous layer was
pipetted off and the precipitate was discarded. 0.5 mL of isopropanol was added, mixed slowly,
and incubated at room temperature for 10 minutes. The mixture was then centrifuged at 12,000 x
g for 10 minutes at 40C. The supernatant was poured off the precipitate and washed with 1 mL
75% ethanol, inverted, and centrifuged at 7,000 x g for 5 minutes at 4oC. The supernatant was
poured off again and the excess ethanol was blotted. The RNA pellet was dried for 5-10 minutes
in the fume hood, dissolved in 25 µL of RNase-free water, and stored at -80oC.
RT-PCR (Reverse Transcription-Polymerase Chain Reaction)
RT-PCR was performed following instructions for the Qiagen One-Step RT-PCR kit.
Primers for aromatase, the RNA of interest, and β-actin, which acted as a loading control, were
used (Table 2; Fenske and Segner, 2004). β-actin was used as the loading control because β-actin
RNA levels stay constant through atrazine exposure, as actin is a cellular cytoskeletal element
that is separate from mechanisms in which atrazine and aromatase are involved. In this way, the
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β-actin RNA acted as a baseline amount of RNA for comparison with the aromatase RNA.
Additionally, the detection band for β-actin is 150 bp, while the band for aromatase is 330 bp, so
a distinction can be made between the two (Mater Methods, 2012; Alberts et al., 1998; Fenske
and Segner, 2004). All four primers were added to non-exposed RNA and exposed RNA.
Primer
Sequence (5’ 3’)
Aromatase forward
CTCCAGCACTCGCATCTGTC
Aromatase reverse
CAGCAGAGCCACCAGAATAGAC
β-actin forward
GCCAACAGAGAGAAGATGACACAG
β-actin reverse
CAGGAAGGAAGGCTGGAAGAG
Table 2. Aromatase and β-actin primers used (Fenske and Segner, 2004).
Gel Electrophoresis
1.5% agarose gel in 0.5 x TBE buffer (Dawson, Molecular Biology Protocols; 89mM
Tris, 89mM boric acid, 2mM EDTA, pH 8) was run for 1 hour at 75 V for Trial 1, and 1 hour
and 45 minutes at 75 V for the Trial 2. Clearest results were seen after about an hour to two
hours at 75 V. A 100 bp ladder was utilized (Qiagen, 210210).
Trial One tested non-exposed RNA and exposed RNA with and without 5Q (Q-solution,
5x concentrated) against known RNA. 5Q is a Qiagen product that changes the melting
behaviors of nucleic acids to increase amplification of low-yield RNA, and was used to
determine if results would differ with or without 5Q (Qiagen, 2015). The known RNA in this
experiment was from wild type (WT) zebrafish embryos that had previously been shown to
produce results in RT-PCR and gel electrophoresis experiments. In initial trials, this RNA acted
as a positive control (in that it was expected to produce RT-PCR products) to improve the
technique, as the first few trials did not show any RT-PCR products. After the method was
improved to obtain readable results, this same RNA was used as a negative control in Trial one
against aromatase expression, as WT zebrafish embryos are not sexually differentiated yet, so
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they were not expected to express aromatase. Trial two was conducted to confirm the results of
the first trial, and tested non-exposed RNA and exposed RNA against the known WT embryo
RNA (negative control).
Results
Sperm Isolation
Sperm counts were unable to be performed effectively due to the fact that consistent
amounts of sperm could not be extracted from each fish. Though a method of moving the sperm
onto the grid of the hemocytometer was devised, the actual sperm count was meaningless as
there was a different initial amount of sperm for each fish. In initial observations, exposed sperm
and non-exposed sperm did not appear to differ morphologically.
Gel Electrophoresis
Trial one was run with a negative control (RNA from WT zebrafish embryos), nonexposed RNA with and without 5Q, and exposed RNA with and without 5Q. This was the
clearest trial, as two distinct bands can be seen for the non-exposed and exposed RNA. The nonexposed sample has two bands, one at aromatase and one at β-actin. The negative control,
exposed with 5Q, and exposed without 5Q samples showed β-actin bands (Figure 4).
Figure 4. Results for the negative
control, non-exposed with and without
5Q, and exposed with and without 5Q.
1.5% agarose gel ran for 1 hour at 75
V. Aromatase and β-actin labels
demarcate the location of expected
bands for the enzyme and protein,
respectively.
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Trial two tested the negative control, non-exposed, and exposed RNA to confirm the
results from trial one. The gel was not as clear during this trial, though bands at aromatase can be
seen in the non-exposed lane, as was seen in the previous trial (Figure 5). The other lanes for the
negative control and exposed samples showed β-actin bands, confirming the results seen in the
previous trial. 5Q was not utilized in this trial because the previous trial did not show any
difference in visualization if 5Q was present or not.
Figure 5. Negative control, nonexposed, and exposed run in a 1.5%
agarose gel at 75 V for 1 hour, 45
minutes. Aromatase and β-actin labels
demarcate the location of expected
bands for the enzyme and protein,
respectively.
Discussion
Sperm Isolation
A method of reliably testing sperm counts was not determined, though it was found that
using Hank’s solution is a valid way to move the sperm onto the hemocytometer grid. With
improvements in the protocol, conducting sperm count tests would be a valuable parameter in
testing the effect of atrazine exposure on male organisms. Expected results for such a protocol
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would be a decreased sperm count for the exposed males when compared to the non-exposed
males, as atrazine is hypothesized to increase aromatase expression or activity, thus, increasing
the levels of estrogen and decreasing the levels of testosterone.
Gel Electrophoresis
Results for the gel electrophoresis did not support the hypothesis: the non-exposed group
had bands indicating aromatase expression, while the exposed group did not. Expected results for
aromatase expression were that the exposed samples would show an increased expression of
aromatase when compared to the non-exposed group of zebrafish.
Experimental Error
Because the results did not support the hypothesis, experimental errors must be
considered. The non-exposed β-actin bands in both trials one and two are less intense than the
exposed and negative control bands, which indicates a lower volume of RNA in the non-exposed
sample. This problem could have been mitigated by adding a greater volume of RNA to the nonexposed lanes in trials one and two. However, because two other trials were performed with the
control RNA before any meaningful results were obtained, there was not a large enough volume
of control RNA for this issue to be addressed. However, this issue could also be because the nonexposed RNA was of less integrity than the exposed RNA. Due to the multiple trials and loss of
control RNA, this study could not be duplicated to confirm the results past reasonable doubt.
In an ideal situation, this entire experiment would have been repeated to confirm the
results, as the possibility that the non-exposed RNA and exposed RNA were mixed during the
testes or RNA isolation exists, however unlikely. However, due to time constraints, such a
duplication was not possible.
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Other Explanations
Aside from experimental errors, there are other reasons to explain why the results of this
study directly refute both the study’s hypothesis and the aromatase hypothesis. One such
explanation is that 70 ppb atrazine is too low of a concentration to see results in zebrafish,
though this conclusion is not supported by a previous study. It is also possible that a 9 day
exposure period is not long enough, though this conclusion is refuted by the same previous study
(Suzawa and Ingraham, 2008). The most likely explanation for the results of this experiment lie
in the distinction between aromatase expression and activity. This study only analyzed aromatase
expression by measuring the amount of RNA, but it may be that atrazine actually effects the
aromatase activity, which this study was unable to test. However, it should be noted that this
explanation does not entirely elucidate why aromatase expression was only seen in the control
organisms for this particular study.
Further studies to clarify the effects of aromatase on zebrafish would be to measure the
activity and expression, respectively, of NR5A receptors and the zycp19a1 gene between a nonexposed and an atrazine-exposed group. This proposed experiment would examine the earlier
stages of the aromatase pathway and may elucidate the exact mechanism by which atrazine
increases aromatase expression or activity. Another important further study would be to compare
aromatase expression and atrazine exposure, much in the same way this study did, with
aromatase activity and atrazine exposure, which can be done via an enzyme activity assay. In
this, the question of if aromatase activity or expression is altered with atrazine exposure may be
answered, which would bring the scientific community closer to an understanding of how
atrazine effects the aromatase mechanism.
Schmaus 20
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