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In ovo inhibition of steroid metabolism by
bisphenol-A as a potential mechanism
of endocrine disruption
rspb.royalsocietypublishing.org
Sandrine G. Clairardin, Ryan T. Paitz and Rachel M. Bowden
School of Biological Sciences, Illinois State University, Campus Box 4120, Normal, IL 61790-4120, USA
Research
Cite this article: Clairardin SG, Paitz RT,
Bowden RM. 2013 In ovo inhibition of steroid
metabolism by bisphenol-A as a potential
mechanism of endocrine disruption. Proc R
Soc B 280: 20131773.
http://dx.doi.org/10.1098/rspb.2013.1773
Received: 9 July 2013
Accepted: 7 August 2013
Subject Areas:
physiology, developmental biology
Keywords:
oestradiol, oestrone sulfate, steroid metabolism
Author for correspondence:
Rachel M. Bowden
e-mail: [email protected]
Electronic supplementary material is available
at http://dx.doi.org/10.1098/rspb.2013.1773 or
via http://rspb.royalsocietypublishing.org.
During embryonic development, endogenous signals, for example steroid
hormones, and exogenous signals, for example endocrine disrupting chemicals (EDCs), have the capacity to produce phenotypic effects that persist
into adulthood. As the actions of steroids are mediated through the binding
of steroid receptors, most studies of EDCs have assumed that they too
elicit their effects by binding steroid receptors. We tested an alternative
hypothesis, namely that EDCs elicit their effects during embryonic development by disrupting the metabolism of maternally derived steroids, thereby
allowing maternally derived steroids to bind steroid receptors and elicit
effects. Specifically, we examined the ability of the EDC, bisphenol-A
(BPA) to inhibit the normal metabolism of oestradiol during the first nine
days of embryonic development in the red-eared slider turtle (Trachemys
scripta). We found that, when BPA was present, oestrogen metabolism was
inhibited when compared to control eggs. In particular, the formation
of oestrone sulfate was blocked in BPA-treated eggs. We postulate that the
oestrogenic effects of EDCs may be driven, at least in part, by inappropriate
oestrogen signalling. The retention of oestrogens at points of development
when they would normally be metabolized to inactive forms might also
help explain low-dose effects frequently reported for EDCs.
1. Introduction
It has long been recognized that the endocrine environment experienced by
developing vertebrate embryos can produce long-lasting phenotypic effects
(reviewed in [1]). Many of these effects are unique to the embryonic period
such that similar endocrine signals later in life are not capable of producing
the same effect. The heightened sensitivity of embryos to the endocrine environment has led to a great deal of research focused on understanding how early
endocrine signals influence development. During embryonic development,
vertebrate embryos are exposed to steroids of embryonic and maternal origin.
Because one of the main functions of oestradiol is to regulate reproduction in
females [2], there is an inherent link between maternal oestradiol and offspring
development. For many vertebrates, maternal oestradiol levels increase during
reproduction [2]. In placental vertebrates, this results in embryos developing
within a maternal environment that contains elevated oestradiol concentrations
[3], whereas in oviparous vertebrates, this results in the presence of oestradiol,
and other lipophilic steroids, in the yolk at oviposition [4].
Research on the early endocrine environment has taken on additional
importance in light of the fact that a large number of environmental and
man-made chemicals often interfere with normal endocrine signalling, and produce long-lasting phenotypic effects, including effects similar to those observed
following oestrogen exposure [5]. These endocrine disrupting chemicals (EDCs)
are comprised of a structurally diverse group of compounds that are also
known as ‘xenoestrogens’ [6], ‘endocrine disrupting chemicals with oestrogenic
activity (EEDCs)’ [7], ‘oestrogen mimics’ [8] and ‘environmental oestrogens’ [9].
A major focus of ongoing research is to determine how such a structurally
diverse group of chemicals could produce effects similar to such a tightly regulated ligand, for example oestradiol, with an emphasis on the ability of EDCs to
& 2013 The Author(s) Published by the Royal Society. All rights reserved.
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sulfonation of oestradiol in vitro [18]. These characteristics
make BPA an ideal compound for investigating the potential
for EDCs to produce oestrogenic effects by inhibiting the
natural metabolism of maternal oestradiol.
2. Material and methods
(a) Egg collection
(b) Effect of bisphenol-A on in ovo oestradiol
metabolism
To determine the effect of BPA on the early in ovo decline of oestradiol in T. scripta, seven clutches were collected and eggs from
each clutch were randomly allocated to one of two treatments:
control eggs ¼ 0.1 mg oestradiol in 5 ml vehicle (70% ethanol),
or BPA-treated eggs ¼ 0.1 mg oestradiol plus 40 mg BPA in 5 ml
vehicle. For oestradiol, the dose was selected to elevate levels
of oestradiol in the early season clutches to levels found in late
season clutches [25]. For BPA, we performed a pilot study
(data not shown), basing our initial doses on those used in
Stoker et al. [48]; we determined that 40 mg of BPA, given on
day 0 of incubation, did not result in significant egg mortality
in T. scripta. Treatment occurred within 24 h of the start of chalking, a sign that the egg contains a viable embryo that has begun
to develop. For this study, only clutches (n ¼ 5) in which all eggs
began chalking within the first 24 h after laying were used to
control timing of treatment. Within each treatment, eggs from
each clutch were randomly assigned to a sampling day either 1,
3, 6 or 9 days post-treatment. Eggs (n ¼ 4–5 per sampling day
and treatment) were incubated at 278C in moist vermiculite
(approx. 2150 kPa) until sampling, at which point the eggs were
frozen at 2208C until analysis. For analysis, each egg was dissected
and components (yolk and albumen/fluid) were separated and
weighed to the nearest 0.01 g.
Levels of oestrogens in the yolk were measured using radioimmunoassay (RIA) (modified from [49,50]). Oestradiol and
oestrone were measured in independent assays with slightly
modified extraction techniques. For oestradiol, 50 mg of yolk
was diluted and homogenized in 1 ml distilled water. Two thousand cpm of tritiated oestradiol (Perkin Elmer, Waltham, MA,
USA) was added to each sample to allow for the calculation of
per cent recovery. Steroids were extracted with ether (30 : 70,
petroleum ether : diethyl ether) producing two phases: an
aqueous phase and an organic phase, which contain the watersoluble and lipid-soluble molecules, respectively. The organic
phase, containing any free steroids present in the sample,
was then dried down under nitrogen gas, resuspended in 1 ml
of 90% ethanol, vortexed and held at 2208C overnight. The
samples were then centrifuged at 2000 r.p.m. for 5 min at 08C
to remove neutral lipids and proteins. The supernatant was
Proc R Soc B 280: 20131773
In this species, there is consistent variation in levels of maternal
oestradiol with concentrations in the eggs from a female’s second
clutch of the nesting season being approximately 10 higher
than those from her first clutch of the season [25]. In both first
and second clutches, these concentrations decline to undetectable
levels early in development [25]. For this study, eggs were collected at Banner Marsh State Fish and Wildlife Area in Fulton
Co., IL, USA [24,25] either from freshly laid nests in the field
or from gravid females that were caught in baited hoop traps
and transported to Illinois State University where egg laying
was induced by injection of oxytocin [46,47]. All adult trapping,
handling and egg collection methods were approved by the
Illinois Department of Natural Resources Illinois (IDNR permit
NH11.2084) and the Illinois State University Institutional
Animal Care and Use Committee (04-2010).
2
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bind to steroid receptors as a potential mechanism for
producing steroid-like effects [5].
Previous research has found that a variety of EDCs can
bind to oestrogen receptors (ERs) [10,11], suggesting that
these EDCs may elicit oestrogenic effects by direct ER activation, thereby increasing ER-dependent gene transcription
[11,12]. This model of EDC action via ER binding assumes
a promiscuous ER-binding site owing to the extreme variation in EDC structure [13]. While it is clear that EDCs can
bind ERs, when binding does occur, only weak, partial agonistic actions are noted [14], such that the activity of an EDC is
often significantly weaker than that of the natural ligand
[15,16]. Together, the evidence suggests that while ER activation by EDCs may play a role in producing oestrogenic
endpoints, it is not likely the sole mechanism for oestrogenic
effects in all tissues. Other possible mechanisms for the oestrogenic effects of EDCs include the induction of oestradiol
production [17] and the inhibition of oestradiol metabolism
[18,19]. Both of these mechanisms would result in elevated
oestradiol levels and do not require a direct interaction
between the EDC and an ER.
Understanding how EDCs influence steroid metabolism in
developing embryos could be especially important because of
the high sensitivity of the embryo to steroid signals paired with
the elevated levels of maternal oestrogens. A key metabolic
pathway in the metabolism of maternal steroids, including
oestradiol, is sulfonation [20] because the primary conjugative
pathway found in adults (glucuronidation) does not fully
develop until after birth [21]. The sulfonation of steroids is carried out by a family of sulfotransferase enzymes that transfer a
sulfonate group to the acceptor steroid and use a universal
sulfonate donor (30 -phosphoadenosine 50 -phosphosulfate or
PAPS) to carry out this reaction [22]. The sulfonation of steroids
typically reduces or eliminates the ability of the steroid to bind
its respective steroid receptor [23], while also decreasing the
hydrophobicity of the steroid to facilitate clearance [21].
Recent work in reptiles [24–27] and birds [28–30] has demonstrated that in ovo metabolism of maternal steroids occurs.
Characterization of this metabolism in the red-eared slider
turtle (Trachemys scripta) demonstrates that oestradiol is converted to a variety of oestrogen sulfates ( primarily oestrone
sulfate) [24,26,27,31] through sulfonation. Interestingly, sulfotransferase enzymes not only can be subjected to inhibition
by EDCs [32,33], but are also vital to the metabolism and clearance of exogenous chemicals such as those that act as EDCs
[34], creating a situation where there is a shared metabolic
fate for both maternal steroids and EDCs. Given that both
endogenous steroids and EDCs use sulfonation for clearance,
it is possible that the presence of an EDC might affect the
normal clearance of oestradiol.
Here, we investigate the effect of the EDC, bisphenol-A
(BPA), on the metabolism of maternal oestradiol in T. scripta.
BPA is a commonly used plasticizer that is one of the most
mass-produced chemicals in the world with an essentially
ubiquitous presence in the environment ([35], reviewed in
[36]). It has been found around the world in human [36,37]
and non-human animals alike, and is known to have oestrogenic effects on developing organisms across vertebrate and
invertebrate taxa (reviewed in [38 –41]). Although BPA has
been shown to bind to both ERa [15,42] and ERb [42], the
binding affinity for BPA is up to 12 500-fold lower than for
oestradiol [43]. Importantly, it has been shown that BPA is
subject to sulfonation [44,45] and can also inhibit the
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oestradiol (ng g–1)
(a)
3.5
3
3.0
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2.5
2.0
1.5
1.0
0.5
0
oestrone (ng g–1)
(b)
oestrone sulfate (ng g–1)
(c)
1.0
0.9
0.8
0.7
0.6
0.5
0.4
0.3
0.2
0.1
0
Proc R Soc B 280: 20131773
poured off, dried down under nitrogen gas and resuspended
in 10% ethyl acetate in isooctane and loaded onto celitepacked columns. For oestrone, 100 mg of yolk was diluted and
homogenized in 200 ml of distilled water and 2000 cpm of tritiated oestone (Perkin Elmer) was added to calculate recoveries.
To precipitate proteins and neutral lipids, 1 ml of methanol
was added and samples were held at 2208C overnight. The
samples were centrifuged at 2000 r.p.m. for 15 min and the
supernatant was decanted. An additional 1 ml of methanol was
added, and the samples were centrifuged again, and the supernatants were combined and dried under nitrogen. The dried
sample was resuspended in 1 ml of distilled water and 2 ml of
diethyl ether was added, producing an aqueous phase and an
organic phase, respectively. The extraction was repeated, the
organic phases combined and dried under nitrogen gas, and
resuspended in 10% ethyl acetate in isooctane loaded onto
celite-packed columns.
Both steroids were eluted with solutions containing progressively higher concentrations of ethyl acetate in isooctane (0, 10, 20
and 40%). Oestrone was isolated with the 20% fraction, whereas
oestradiol was isolated with the 40% fraction. Duplicate aliquots
of each steroid were subjected to RIA using specific antibodies
(oestrone antibody E3135, Sigma, USA; oestradiol antibody
7010, Biogenesis, UK). To quantify endogenous oestrogens, a
standard curve (range: 1.95– 500 pg) was used for comparison.
Oestrone was analysed in a single assay, with an intra-assay
coefficient of variation (CV) of 4%. Oestradiol was analysed
across two assays, with intra-assay CVs of 13 and 20%, and an
inter-assay CV of 23%.
Oestrone sulfate, oestriol sulfate and oestradiol sulfate were
quantified using liquid chromatography – mass spectrometry
(LC/MS/MS). Samples were prepared by vortexing 1 ml of albumen with 3 ml of methanol (HPLC grade), frozen overnight and
centrifuged (2000 r.p.m.) at 08C for 20 min. The supernatant was
poured off and kept at 2208C. A second methanol extraction was
performed on the pellet with 2 ml of methanol following the
same protocol. Supernatants from both rounds were consolidated, dried down under nitrogen gas and resuspended in
5 ml nano-pure water. Water-soluble metabolites were extracted
using Sep-pak cartridges (C18 plus short cartridge, 55– 105 mm
particle size; Waters, Milford, MA, USA). For each sample, two
Sep-pak cartridges were stacked and charged with 5 ml methanol followed by 5 ml nano-pure water. The samples were then
loaded onto the cartridges. Lipid-soluble molecules were eluted
with 5 ml ether followed by elution of the water-soluble metabolites with 5 ml methanol. The methanol eluate was then dried
down under nitrogen gas and metabolites characterized using
LC/MS/MS.
The LC/MS/MS analysis was performed in the Metabolomics Center at University of Illinois at Urbana – Champaign
with a 5500 QTRAP mass spectrometer (AB Sciex, Foster City,
CA, USA), that is equipped with a 1200 Agilent HPLC. ANALYST
(v. 1.5.1, Applied Biosystems) was used for data acquisition and
processing. The HPLC flow rate was set at 0.35 ml min21. HPLC
mobile phases consisted of 0.1% formic acid in water (A) and
0.1% formic acid in acetonitrile (B). The autosampler was kept
at 58C. The injection volume was 5 ml. An Agilent Zorbax
SB-Aq column (3.5 mm, 50 4.6 mm) was used for the separation with the following gradient: 0 –2 min, 80% A; 6 – 10 min,
25% A; 11 – 16.5 min, 80% A. Mass spectrometer was operated
under negative mode electrospray ionization. The electrospray voltage was set to 24500 V, the heater was set at
6008C, the curtain gas was 35, and GS1 and GS2 were 50, 65,
respectively. Quantitative analysis was performed via MRM
where m/z 349.1 ! m/z 269.1 (3-OL-17-one; oestrone sulfate);
m/z 351.1 ! m/z 271.1 (17-b oestradiol; oestradiol sulfate) and
m/z 367.1 ! m/z 287.1 (16-a-17-b-oestriol; oestriol sulfate)
were monitored.
800
700
600
500
400
300
200
100
0
2
4
6
developmental day
8
10
Figure 1. Levels of (a) oestradiol, (b) oestrone and (c) oestrone sulfate in
T. scripta eggs during early embryonic development. Control eggs (0.1 mg
oestradiol, filled symbols, n ¼ 17) had lower levels of oestradiol and oestrone, but higher levels of oestone sulfate than eggs treated with
bisphenol-A (0.1 mg oestradiol þ 40 mg BPA, open symbols, n ¼ 18).
Data represent back-transformed means of log-transformed data +s.e.
(c) Statistical analyses
Levels of oestrone, oestradiol and oestrone sulfate were analysed
by ANOVA in PROC GLM (SAS 9.3 SAS Institute, Cary, NC,
USA). For the ANOVAs, treatment (control or BPA treated),
sampling day (1, 3, 6 or 9) and their interaction were included
as fixed factors in the model. Clutch of origin was included as
a random factor. We ran contrasts to probe any significant interactions. Levels of oestradiol sulfate were low to undetectable,
while oestriol sulfate levels were below detection limits in all
samples; neither was included in the final analyses. To meet
normality assumptions of ANOVA, all steroids were log(10)
transformed prior to analysis.
3. Results
Mean levels of oestradiol, oestrone and oestrone sulfate in
eggs (n ¼ 1 per clutch) sampled prior to any treatment were
2.7, 0.10 and 5.6 ng g21, respectively. Levels of oestradiol in
the yolk were significantly affected by treatment, with eggs
from the BPA treatment group having elevated levels of
yolk oestradiol relative to those from the control treatment
(F1,1 ¼ 11.10, p ¼ 0.003; figure 1a). There were no significant
effects of either day (F1,3 ¼ 2.50, p ¼ 0.085), or the interaction
of treatment day (F1,3 ¼ 0.19, p ¼ 0.90) on yolk oestradiol.
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E1
E1S
BPA treated
E2
E1
E1S
However, we did detect a significant clutch effect on yolk
oestradiol levels (F1,4 ¼ 18.16, p , 0.0001). Levels of oestrone
in the yolk were also significantly affected by treatment, with
eggs from the BPA treatment group having elevated levels of
yolk oestrone relative to those from the control treatment
(F1,1 ¼ 6.86, p ¼ 0.015; figure 1b). Yolk oestrone levels varied
by sampling day (F1,3 ¼ 12.06, p , 0.0001), but there was no
significant interaction detected (F1,3 ¼ 0.09, p ¼ 0.97). Levels
of oestrone sulfate in the albumen varied significantly by treatment group (F1,1 ¼ 19.30, p ¼ 0.0002) and by day (F1,3 ¼ 54.55,
p , 0.0001). There was a significant treatment day interaction (F1,3 ¼ 4.94, p ¼ 0.0086; figure 1c). Probing this
interaction, we found that the treatment groups varied on
day 1 ( p , 0.0001) and day 6 ( p ¼ 0.006), but not on days 3
and 9 ( p . 0.05). There were no clutch effects detected for
either oestrone or oestrone sulfate. Data used in these analyses
are available as the electronic supplementary material.
4. Discussion
Here, we demonstrate that BPA alters the metabolism of
oestrogens in ovo. We found that levels of both oestradiol and
oestrone were higher, whereas levels of oestrone sulfate were
lower, in eggs treated with BPA when compared with the control
eggs. The low levels of oestrone sulfate paired with the elevated
levels of free oestrogens indicate that oestrogen sulfotransferase
activity is altered in the presence of BPA in T. scripta (figure 2).
Because, however, there are several metabolic conversions
required to produce oestrone sulfate from oestradiol, it is also
possible that BPA could be affecting other steps in this process.
For example, BPA could alter the conversion of oestradiol to oestrone instead of affecting the sulfonation of oestrone itself, thus
producing the lower oestrone sulfate levels observed in the
BPA-treated group. If the conversion from oestradiol to oestrone
was the main pathway inhibited, we would have expected to see
decreased levels of oestrone in the BPA-treated eggs. In fact, we
found just the opposite pattern, indicating that BPA is inhibiting
the conversion of oestrone to oestrone sulfate. Our data support
the hypothesis that EDCs, for instance BPA, can produce their
effects by increasing the bioavailability of active steroids
during critical periods of development by effectively interfering
with the metabolism of maternal steroids.
The ability of EDCs to inhibit the sulfonation of oestrogens
has been demonstrated in a number of in vitro systems ([18,19],
reviewed in [51]), but to our knowledge, this is the first time it
(a) EDC effects in vertebrates
EDCs have been shown to elicit a wide variety of effects from
the ‘classic’ feminizing effects of DDT on gull embryos [52]
to more recent work demonstrating transgenerational epigenetic effects (reviewed in [53]). Interestingly, many effects are
observed when embryos are exposed to low doses of the compounds, and the effects of the EDC can be influenced by
endogenous steroid levels. For example, fetal mice with higher
levels of endogenous oestradiol responded to exposure to a
low dose of BPA in utero, whereas those with low endogenous
oestradiol levels did not [54]. It is possible, then, that even
EDC exposures considered to be ‘low doses’, may be high
enough relative to the available enzyme to produce altered
steroid metabolism, resulting in the inhibition of sulfotransferase activity toward endogenous and exogenous compounds
alike, which may account for the low-dose effects often observed
(see [41] for discussion of dose–response curves for BPA).
This is the first study, to our knowledge, that demonstrates
alteration of steroid metabolism by an EDC in ovo. While in vitro
data provides evidence for the potential of an EDC to inhibit
steroid metabolism, the in vitro approach may lack the feedback
and compensatory mechanisms that may otherwise prevent
alteration in vivo. By demonstrating alteration in a closed and
complete developmental environment, we provide support
for altered steroid metabolism as a mechanism of endocrine
disruption in the whole organism. This altered steroid metabolism could produce inappropriate signalling by endogenous
compounds, potentially producing or contributing to the
known effects of EDCs on tissue differentiation. Although we
have only discussed oestrogens here, the implications of this
research may extend to any endogenous signalling molecules
that use similarly promiscuous conjugation pathways. The
relative promiscuity of conjugative enzymes as compared to
steroid receptors suggests that altered steroid metabolism has
the potential to be a more universal mechanism across
4
Proc R Soc B 280: 20131773
Figure 2. Schematic of how steroid metabolism is altered during the first
nine days of embryonic development in T. scripta when eggs are exposed
to the endocrine disruptor bisphenol-A. In the control eggs, oestradiol (E2)
is converted to oestrone (E1), and then to oestrone sulfate (E1S) which is present at high levels in the egg. Eggs treated with BPA maintained higher
levels of both E2 and E1, with little to no E1S being formed. Our data indicate
that BPA interferes with the metabolic conversion of E1 to E1S in ovo.
has been shown that the sulfonation of maternal oestrogens is
subject to inhibition by an EDC in ovo. In both placental and
oviparous vertebrates, the sulfonation of steroids is thought
to serve as a buffer that protects the embryo from being
exposed to the high levels of maternal oestrogens [20,24].
The presence of increased levels of free oestradiol and oestrone
in the yolks of BPA-treated eggs suggests that inhibiting sulfotranferase activity may lead to increased levels of active steroid
that is potentially available for signalling within the egg.
Demonstrating that the sulfonation of maternal oestrogens
is subject to inhibition by BPA has important implications for
developing vertebrate embryos. Previous work has demonstrated that EDCs can be metabolized through the same
enzymatic pathways as steroids, including sulfonation [34],
and our data indicate that the presence of BPA results in the
inhibition of oestrogen metabolism. At present, it is unclear if
the observed inhibition is the result of competitive inhibition
of BPA for access to the sulfotransferase enzyme, PAPS
depletion, or if non-competitive or allosteric inhibition may
be taking place. Regardless, if embryos are simultaneously
exposed to both maternal oestrogens and exogenous compounds that can be metabolized through the sulfotransferase
pathway, it appears that they may not be able to adequately
remove all active compounds in a timely manner, ultimately
producing effects on the offspring’s phenotype.
rspb.royalsocietypublishing.org
control
E2
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structurally varied EDCs and to better account for low-dose
effects than the traditional receptor-mediated mechanism.
bolomics Center for performing the LC/MS/MS analysis, Steve
References
1.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
action of endogenous and environmental estrogens.
Pflugers Arch. 449, 335–343. (doi:10.1007/
s00424-004-1343-9)
Katzenellenbogen JA. 1995 The structural
pervasiveness of estrogenic activity. Environ. Health
Perspect. 103, 99 –101. (doi:10.2307/3432517)
Mueller SO. 2004 Xenoestrogens: mechanisms of
action and detection methods. Anal. Bioanal. Chem.
378, 582 –587. (doi:10.1007/s00216-003-2238-x)
Gould JC, Leonard LS, Maness SC, Wagner BL,
Conner K, Zacharewski T, Safe S, McDonnell DP,
Gaido KW. 1998 Bisphenol A interacts with the
estrogen receptor a in a distinct manner from
estradiol. Mol. Cell. Endocrinol. 142, 203–214.
(doi:10.1016/S0303-7207(98)00084-7)
Matsushima A, Shimohigashi Y. 2008 A strategy to
explore the target receptor of endocrine disruptors:
estrogen-related receptor g (ERR g) as a genuine
acceptor of bisphenol A. In Proc. 6th World Congress on
Alternatives & Animal Use in the Life Sciences, 21–25
August 2007, Tokyo, Japan. AATEX 14, 495–497.
Chung E, Genco MC, Megrelis L, Ruderman JV. 2011
Effects of bisphenol A and triclocarban on brainspecific expression of aromatase in early zebrafish
embryos. Proc. Natl Acad. Sci. USA 108, 17 732–
17 737. (doi:10.1073/pnas.1115187108)
Kester MHA et al. 2002 Potent inhibition of estrogen
sulfotransferase by hydroxylated metabolites of
polyhalogenated aromatic hydrocarbons reveals
alternative mechanism for estrogenic activity of
endocrine disrupters. J. Clin. Endocrinol. Metab. 87,
1142 –1150. (doi:10.1210/jc.87.3.1142)
Harris RM, Wood DM, Bottomley L, Blagg S, Owen
K, Hughes PJ, Waring RH, Kirk CJ. 2004
Phytoestrogens are potent inhibitors of estrogen
sulfation: implications for breast cancer risk and
treatment. J. Clin. Endocrinol. Metab. 89,
1779 –1787. (doi:10.1210/jc.2003-031631).
Diczfalusy E. 1964 Endocrine functions of human
fetoplacental unit. Fed. Proc. 23, 791 –798.
Dutton GJ. 1978 Developmental aspects of drug
conjugation, with special reference to
glucuronidation. Annu. Rev. Pharmacol. Toxicol. 18,
17 –35. (doi:10.1146/annurev.pa.18.040178.
000313)
Strott CA. 2002 Sulfonation and molecular action.
Endocr. Rev. 23, 703–732. (doi:10.1210/er.
2001.0040)
Pasqualini JR, Gelly C, Lecerf F. 1986 Estrogen
sulfates: biological and ultrastructural responses and
metabolism in MCF-7 human breast cancer cells.
Breast Cancer Res. Treat. 8, 233 –240. (doi:10.1007/
bf01807336)
24. Paitz RT, Bowden RM. 2008 A proposed role of the
sulfotransferase/sulfatase pathway in modulating
yolk steroid effects. Integr. Comp. Biol. 48,
419–427. (doi:10.1093/icb/icn034)
25. Paitz RT, Bowden RM. 2009 Rapid decline in the
concentrations of three yolk steroids during
development: is it embryonic regulation? Gen.
Comp. Endocrinol. 161, 246–251. (doi:10.1016/j.
ygcen.2009.01.018)
26. Paitz RT, Bowden RM. 2011 Biological activity of
oestradiol sulphate in an oviparous amniote:
implications for maternal steroid effects.
Proc. R. Soc. B 278, 2005 –2010. (doi:10.1098/
rspb.2010.2128)
27. Paitz RT, Sawa AR, Bowden RM. 2012 Characterizing
the metabolism and movement of yolk oestradiol
during embryonic development in the redeared slider (Trachemys scripta). Gen. Comp.
Endocrinol. 176, 507–512. (doi:10.1016/j.ygcen.
2011.10.009)
28. von Engelhardt N, Henriksen R, Groothuis TGG. 2009
Steroids in chicken egg yolk: metabolism and
uptake during early embryonic development. Gen.
Comp. Endocrinol. 163, 175–183. (doi:10.1016/j.
ygcen.2009.04.004)
29. Paitz RT, Bowden RM, Casto JM. 2011 Embryonic
modulation of maternal steroids in European
starlings (Sturnus vulgaris). Proc. R. Soc. B 278,
99– 106. (doi:10.1098/rspb.2010.0813)
30. Paitz RT, Casto JM. 2012 The decline in yolk
progesterone concentrations during incubation is
dependent on embryonic development in the
European starling. Gen. Comp. Endocrinol. 176,
415–419. (doi:10.1016/j.ygcen.2011.12.014)
31. Paitz RT, Bowden RM. 2013 The sulfonation of maternal
steroids is a conserved metabolic pathway in
vertebrates. Integr. Comp. Biol. (doi:10.1093/icb/ict027)
32. Ekuase EJ, Liu Y, Lehmler HJ, Robertson LW, Duffel
MW. 2011 Structure–activity relationships for
hydroxylated polychlorinated biphenyls as inhibitors
of the sulfation of dehydroepiandrosterone
catalyzed by human hydroxysteroid sulfotransferase
SULT2A1. Chem. Res. Toxicol. 24, 1720–1728.
(doi:10.1021/tx200260h)
33. Wang LQ, Lehmler HJ, Robertson LW, James MO.
2006 Polychlorobiphenylols are selective inhibitors
of human phenol sulfotransferase 1A1 with
4-nitrophenol as a substrate. Chem. Biol. Interact.
159, 235–246. (doi:10.1016/j.cbi.2005.12.004)
34. Xu CJ, Li CYT, Kong ANT. 2005 Induction of phase I,
II and III drug metabolism/transport by xenobiotics.
Arch. Pharm. Res. 28, 249–268. (doi:10.1007/
BF02977789)
Proc R Soc B 280: 20131773
Arnold AP. 2009 The organizational –activational
hypothesis as the foundation for a unified theory of
sexual differentiation of all mammalian tissues.
Horm. Behav. 55, 570–578. (doi:10.1016/j.yhbeh.
2009.03.011)
2. Norris DO. 2007 Vertebrate endocrinology, 4th edn.
San Diego, CA: Academic Press.
3. Conley AJ, Mason JI. 1990 Placental steroidhormones. Baillière’s Clin. Endocrinol. Metab. 4,
249–272. (doi:10.1016/s0950-351x(05)80050-3)
4. Carere C, Blathazart J. 2007 Sexual versus individual
differentiation: the controversial role of avian
maternal hormones. Trends Endocrinol. Metab. 18,
73 –80. (doi:10.1016/j.tem.2007.01.003)
5. Zoeller RT, Brown TR, Doan LL, Gore AC, Skakkebaek
NE, Soto AM, Woodruff TJ, vom Saal FS. 2012
Endocrine-disrupting chemicals and public health
protection: a statement of principles from The
Endocrine Society. Endocrinology 153, 4097 –4110.
(doi:10.1210/en.2012-1422)
6. Gore AC. 2008 Developmental programming and
endocrine disruptor effects on reproductive
neuroendocrine systems. Front. Neuroendocrinol. 29,
358–374. (doi:10.1016/j.yfrne.2008.02.002)
7. Welshons WV, Thayer KA, Judy BM, Taylor JA,
Curran EM, vom Saal FS. 2003 Large effects from
small exposures. I. Mechanisms for endocrinedisrupting chemicals with estrogenic activity.
Environ. Health Perspect. 111, 994–1006. (doi:10.
1289/ehp.5494)
8. Schug TT, Janesick A, Blumberg B, Heindel JJ. 2011
Endocrine disrupting chemicals and disease
susceptibility. J. Steroid Biochem. Mol. Biol. 127,
204–215. (doi:10.1016/j.jsbmb.2011.08.007)
9. Steinmetz R, Brown NG, Allen DL, Bigsby RM,
BenJonathan N. 1997 The environmental estrogen
bisphenol a stimulates prolactin release in vitro and
in vivo. Endocrinology 138, 1780 –1786. (doi:10.
1210/en.138.5.1780)
10. Kuiper G, Carlson B, Grandien K, Enmark E,
Haggblad J, Nilsson S, Gustafsson J. 1997
Comparison of ligand binding specificity and
transcript tissues distribution of estrogen receptors
a and b. Endocrinology 138, 863–870. (doi:10.
1210/en.138.3.863)
11. Kuiper G, Lemmen JG, Carlsson B, Corton JC, Safe
SH, van der Saag PT, van der Burg P, Gustafsson JA.
1998 Interaction of estrogenic chemicals and
phytoestrogens with estrogen receptor beta.
Endocrinology 139, 4252 –4263. (doi:10.1210/en.
139.10.4252)
12. Nadal A, Alonso-Magdalena P, Ripoll C, Fuentas E.
2005 Disentangling the molecular mechanisms of
5
rspb.royalsocietypublishing.org
Acknowledgements. We thank Zhong Li at the University of Illinois Meta-
Juliano for statistical advice, the Illinois Department of Natural
Resources for access to Banner Marsh, and Laura Zimmerman and
Justin Hicke for assistance in the field.
Funding statement. This work was supported by an EPA STAR
Fellowship to S.G.C., and NSF IOS-0952840 to R.M.B.
Downloaded from http://rspb.royalsocietypublishing.org/ on June 15, 2017
48. Stoker C, Rodriguez H, Ramos JG, Sirosky P, Larriera
A, Luque EH, Muñoz-de-Toro M. 2003 Sex reversal
effects on Caiman latirostris exposed to
environmentally relevant doses of the xenoestrogen
bisphenol A. Gen. Comp. Endocrinol. 133, 287 –296.
(doi:10.1016/S0016-6480(03)00199-0)
49. Wingfield JC, Farner DS. 1975 The determination of
five steroids in avian plasma by radioimmunoassay
and competitive protein binding. Steroids 26,
311–327. (doi:10.1016/0039-128X(75)90077-X)
50. Bowden RM, Ewert MA, Nelson CE. 2000
Environmental sex determination in a reptile varies
seasonally and with yolk hormones. Proc. R. Soc. Lond.
B 267, 1745–1749. (doi:10.1098/rspb.2000.1205)
51. Wang LQ, James MO. 2006 Inhibition of
sulfotransferases by xenobiotics. Curr. Drug Metab.
7, 83 –104. (doi:10.2174/138920006774832596)
52. Fry DM, Toone CK. 1981 DDT-induced feminization
of gull embryos. Science 213, 922–924. (doi:10.
1126/science.7256288)
53. Anway MD, Skinner MK. 2006 Epigenetic
transgenerational actions of endocrine disruptors.
Endocrinology 147, S43–S49. (doi:10.1210/en.
2005-1058)
54. Howdeshell KL, vom Saal FS. 2000 Developmental
exposure to bisphenol A: interaction with
endogenous estradiol during pregnancy in mice.
Am. Zool. 40, 429–437. (doi:10.1668/00031569(2000)040[0429:DETBAI]2.0.CO;2)
6
Proc R Soc B 280: 20131773
41. Vandenberg LN, Maffini MV, Sonnenschein C, Rubin
BS, Soto AM. 2009 Bisphenol-A and the great
divide: a review of controversies in the field of
endocrine disruption. Endocr. Rev. 30, 75 –95.
(doi:10.1210/er.2008-0021)
42. Matthews JB, Twomey K, Zacharewski TR. 2001
In vitro and in vivo interactions of bisphenol A and
its metabolite, bisphenol A glucuronide, with
estrogen receptors a and b. Chem. Res. Toxicol. 14,
149 –157. (doi:10.1021/tx0001833)
43. U.S. EPA. 2005 A cross-species mode of action
information assessment: a case study of bisphenol-A
(BPA) (EPA/600/R-50/044F). Washington, DC: US
Environmental Protection Agency.
44. Hanoika N, Takeda Y, Tanaka-Kagawa T, Hayashi K,
Jinno H, Narimatsu S. 2008 Interaction of bisphenol
a with human UDP-glucuronosyltransferase 1a6
enzyme. Environ. Toxicol. 23, 407 –412. (doi:10.
1002/tox.20345)
45. Fini J-B, Dolo L, Cravedi J-P, Demeneix B, Zalko D.
2009 Metabolism of the endocrine disruptor BPA by
Xenopus laevis tadpoles. Ann. NY Acad. Sci. 1163,
394 –397. (doi:10.1111/j.1749-6632.2008.03655.x)
46. Ewert MA, Legler JM. 1978 Hormonal induction of
oviposition in turtles. Herpetologica 34, 314– 318.
47. Les HL, Paitz RT, Bowden RM. 2007 Experimental
test of the effects of fluctuating incubation
temperatures on hatchling phenotype. J. Exp. Zool.
307A, 274 –280. (doi:10.1002/jez.374)
rspb.royalsocietypublishing.org
35. Talsness CE, Andrade AJM, Kuriyama SN, Taylor JA,
vom Saal FS. 2009 Components of plastic:
experimental studies in animals and relevance
for human health. Phil. Trans. R. Soc. B 364,
2079–2096. (doi:10.1098/rstb.2008.0281)
36. Vandenberg LN, Hauser R, Marcus M, Olea N,
Welshons WV. 2007 Human exposure to bisphenol A
(BPA). Reprod. Toxicol. 24, 139–177. (doi:10.1016/
j.reprotox.2007.07.010)
37. Vandenberg LN, Chahoud I, Heindel J,
Padmanabhan V, Paumgartten FJR, Schoenfelder G.
2010 Urinary circulating and tissue biomonitoring
studies indicate widespread exposure to bisphenol
A. Environ. Health Perspect. 118, 1055 –1070.
(doi:10.1289/ehp.0901716)
38. McLachlan JA. 2001 Environmental signaling: what
embryos and evolution teach us about endocrine
disrupting chemicals. Endocr. Rev. 22, 319– 341.
(doi:10.1210/er.22.3.319)
39. Welshons WV, Nagel SC, vom Saal FS. 2006 Large
effects from small exposures. III. Endocrine
mechanisms mediating effects of bisphenol A at
levels of human exposure. Endocrinology 147,
S56–S69. (doi:10.1210/en.2005-1159)
40. Crain DA, Eriksen M, Iguchi T, Jobling S, Laufer H,
LeBlanc GA, Guillette LJ. 2007 An ecological
assessment of bisphenol-A: evidence from
comparative biology. Reprod. Toxicol. 24, 225–239.
(doi:10.1016/j.reprotox.2007.05.008)