Effects of oocyte bisphenol A exposure on aspects of oocyte

Effects of oocyte bisphenol A exposure on aspects of oocyte maturation and
early embryo development in Bos taurus
by
Jacqueline Christine Ferris
A Thesis
presented to
The University of Guelph
In partial fulfilment of requirements
for the degree of
Doctor of Philosophy
in
Biomedical Sciences
Guelph, Ontario, Canada
© Jacqueline Christine Ferris, May, 2015
ABSTRACT
EFFECTS OF OOCYTE BISHPENOL A EXPOSURE ON ASPECTS OF OOCYTE
MATURATION AND EARLY EMBRYO DEVELOPMENT IN BOS TAURUS
Jacqueline Christine Ferris
University of Guelph, 2015
Advisors:
Dr. Allan King
Dr. Neil MacLusky
The microenvironment of the oocyte can influence oocyte quality, competence, and
developmental potential. Alterations in this environment can have negative effects on oocyte
maturation and subsequent embryo development. Oocyte quality, which can determine embryonic
viability, is easily perturbed, thus factors that can alter normal oocyte maturation are a considerable
concern. Bisphenol A (BPA) is an endocrine disrupting chemical that has been found to elicit a
variety of reproductive effects. Exposure to BPA is considered to be ubiquitous and it has been
found in a variety of human samples, including follicular fluid. BPA has previously been found to
disrupt meiosis in mouse and human oocytes, however the embryonic effect in mammals are not
as well documented. In the current study, bovine oocytes were matured in vitro under various
treatment conditions such as no-treatment control (IVM media), vehicle control (0.1% ethanol),
estradiol (2 μg/mL E2), and 15 ng/mL and 30 ng/mL BPA. The mature oocytes or subsequent
embryos were collected for various analyses to determine effects on quality and developmental
potential. Exposure of groups of oocytes to 15 ng/mL and 30 ng/mL BPA resulted in an average
oocyte uptake of 1.69 and 2.48 ng/mL BPA, respectively. Exposure of bovine oocytes in vitro to
30 ng/mL BPA during maturation induced meiotic perturbations as well as poor embryonic
outcomes. Meiosis progression was reduced and abnormal spindle morphology and chromosome
alignment were increased. Under the same treatment conditions, resulting embryos exhibited
decreased embryonic development rates, increased apoptosis, and a skewed sex ratio. Gene
expression in blastocysts were not altered, whereas treatment with 15 ng/mL BPA resulted in
increased expression of CDC2, AURKA, DAZL, TRβ and p53 in MII oocytes relative to that of
the IVM group. This increased p53 expression resulting from the 15 ng/mL BPA group was also
significantly greater than the expression levels resulting from 30 ng/mL BPA. There appeared to
be a slight vehicle effect, with the vehicle group (0.1% ethanol) resulting in significantly increased
expression of AURKA mRNA and non-significant increases in several other genes analyzed. BPA
exposure during oocyte maturation in vitro can therefore, in a dose-dependent way, decrease
oocyte and embryo quality and developmental potential.
iv
DECLARATION OF WORK PERFORMED
I declare that with the exception of the items indicated below, all work reported in the body
of this thesis was performed by me.
Cattle ovaries were collected by Pradeep Balaraju, Steven Huang, and Heather Smale.
Media used for oocyte in vitro maturation, in vitro fertilization, and in vitro culture were prepared
by Liz St. John. Oocytes were collected for real time analysis with the help of Dr. Teresa Mogas,
and the real time qPCR experiments were performed by Kiana Mahboubi.
v
ACKNOWLEDGEMENTS
First and foremost I would like to extend my gratitude to Dr. Allan King, my supervisor
and mentor, who truly made all of this possible. Allan provided me with a perfect balance of
support, encouragement, advice, and ideas, while still allowing me to follow my instincts and work
independently. I have learned a lot working with Allan, and am very fortunate to have had the
opportunity to work under his advisement. I have grown tremendously since starting my PhD and
Allan’s support and confidence in me has played a major role in this development. Thank you
Allan for all you have done to support me throughout this journey!
I would like to thank Dr. Neil MacLusky, my co-advisor who has provided support, advice,
statistical support, and much guidance throughout my time at the University of Guelph. I have
learned a lot from Neil both through my graduate work, as his teaching assistant, as well as from
the conversations that we’ve shared. I consider myself lucky for the time I have been able to work
with him. Thank you Neil for providing me with encouragement and support, and for all the advice
you have given me.
I would like to thank Dr. John Leatherland, who supervised my MSc studies and continued
to advise me as part of my committee for my PhD studies. John took me on as an MSc student
although I had little laboratory experience. He provided me with guidance and support, while
encouraging me to work independently, allowing me to develop skills that prepared me for my
time as a PhD student. He has continued to provide me with support and encouragement and was
instrumental in the revising of my thesis. Thank you John for taking a chance on me and
introducing me to the world of research!
vi
I would sincerely like to thank Dr. Nicholas Bernier, who served as my external committee
member, for all of his support, input, and assistance during my PhD studies. He was always
encouraging and optimistic. Nick helped to revise my thesis, providing valuable insight which
helped me to compile a better thesis. Thank you Nick for all of your support over the years!
I’d like to give a special mention to Dr. Laura Favetta who was a constant source of
guidance, encouragement, and support for me. Laura helped me with design ideas, strategies,
technical guidance, and damage control, helping keep my project on track. Laura was a mentor to
me throughout my PhD studies. Thank you Laura for all the support you have given me, I truly
could not have done this without you!
I would like to thank Dr. Monica Antenos, Lis St. John, Ed Reyes, Allison MacKay, Dr.
Teresa Mogan, Yu Gu, Helen Coates, Tamas Revay, and Kiana Mahboubi for their technical
advice and support. I would like to give a special acknowledgement to Liz for all of her help with
in vitro embryo production, as well as other aspects of my project. I learned a lot from Liz and am
very fortunate to have benefited from her expertise, advice, and technical help. I would also like
to give a special thank you to Monica who has been a constant source of support and advice, and
for her help reviewing a section of this thesis. A big thank you goes to Teresa for helping me with
time-sensitive oocyte collections, and Kiana who conducted the real-time experiments on a tight
schedule. Without Kiana’s help my study period would surely have been extended.
I would like to give a special thank you to members of my former laboratory, Dr. Mao Li
and Lucy Lin. Together they guided me through my first two years in graduate school and provided
me with help whenever it was needed. They helped train me and taught me much of what I know
about working in a laboratory as well as a graduate researcher. They were always very encouraging
vii
and a great source of support and advice. I am lucky to have had them as mentors during my early
graduate career.
This research would not have been possible without the financial support from the Natural
Sciences and Engineering Research Council, and Canada Research Chairs program. I also would
not have been able to conduct my studies without support from the OVC Scholarship program as
well as the support I received from Ontario Graduate Scholarship (OGS).
I have received help from many colleagues both in and out of the laboratory whether it be
technical advice or help, support, encouragement, or their friendship. Thank you to all those who
had a role in my experiences over the last several years with a special thank you to Kayla, Carmon,
Anja, Carolyn, Les, Graham, Allison, Jeff, Stewart, Tim, Nayoung, and Fazl. I would also like to
thank Nathalia and Priscilla for their help and friendship in their time at Guelph.
Outside of school, I owe many thanks to those who have helped me along the way and have
played a major role in the person I have become. I would like to thank my parents for encouraging
my education and teaching me that I can do anything I put my mind to. Thank you for your constant
support and challenging me to be the best person I can be. Thank you to my family and friends
who have encouraged me to be my best self and follow my dreams just by being the amazing
people that they are and for their words of encouragement, love, and support along the way.
I have to give a very special thank you to my husband, Simeon. He has been an incredible
source of support and encouragement. He is always my biggest supporter and helped me
immensely I these final months allowing me to focus on writing, even though he had his own thesis
to complete. I knew I could do this with him by my side. We did it!
viii
TABLE OF CONTENTS
DECLARATION OF WORK PERFORMED ............................................................................... iv
ACKNOWLEDGEMENTS .............................................................................................................v
LIST OF TABLES ......................................................................................................................... xi
LIST OF FIGURES ...................................................................................................................... xii
LIST OF ABBREVIATIONS ........................................................................................................xv
INTRODUCTION ...........................................................................................................................1
LITERATURE REVIEW ................................................................................................................4
Mammalian oogenesis ..................................................................................................................4
Oocyte maturation ....................................................................................................................5
Oocyte competence ..................................................................................................................8
The follicular environment .......................................................................................................9
Environmental influences .......................................................................................................11
Early embryo development ........................................................................................................13
Vulnerability of the embryo ...................................................................................................14
Apoptosis in the preimplantation embryo ..............................................................................16
Embryo sex ratio.....................................................................................................................17
Blastocyst gene expression.........................................................................................................20
Hormone receptors .................................................................................................................22
Embryonic stress, metabolism, and gene expression .............................................................24
Bisphenol A ................................................................................................................................27
Metabolism and concentration ...............................................................................................28
Known effects of BPA............................................................................................................31
BPA and the oocyte ................................................................................................................32
BPA and male reproductive effects ........................................................................................34
BPA and gene expression .......................................................................................................35
BPA and gene expression .......................................................................................................34
Nuclear receptors .................................................................................................................36
Stress and metabolism .........................................................................................................38
RATIONALE, HYPOTHESIS AND OBJECTIVES ....................................................................40
Rationale .................................................................................................................................40
Hypothesis ..............................................................................................................................41
ix
Objectives ...............................................................................................................................41
CHAPTER ONE ............................................................................................................................42
INTRODUCTION ......................................................................................................................43
MATERIALS AND METHODS ...............................................................................................45
Experimental design ...............................................................................................................45
Chemicals ...............................................................................................................................45
Oocyte collection and in vitro oocyte maturation ..................................................................45
Enzyme-linked Immunosorbent Assay (ELISA)....................................................................46
Immunocytochemistry and Imaging .......................................................................................48
Analysis of oocytes.................................................................................................................49
Spindle formation and chromosome alignment analyses .......................................................49
Statistical analyses ..................................................................................................................50
RESULTS...................................................................................................................................51
Oocyte uptake of BPA and IVM media concentrations .........................................................51
Meiosis progression and MII spindle abnormalities ..............................................................51
DISCUSSION ............................................................................................................................60
Oocyte uptake of BPA and IVM media concentrations .........................................................61
Meiosis progression and MII spindle abnormalities ..............................................................63
CHAPTER TWO ...........................................................................................................................68
INTRODUCTION ......................................................................................................................69
MATERIALS AND METHODS ...............................................................................................73
Experimental design ...............................................................................................................73
Chemicals ...............................................................................................................................73
Oocyte collection and in vitro embryo production .................................................................73
Calculation of development rates ...........................................................................................74
Embryo sexing my PCR .........................................................................................................74
TUNEL analysis of apoptosis.................................................................................................75
Statistical analyses ..................................................................................................................78
RESULTS...................................................................................................................................79
Developmental rates, sex ratio, and total cell number of blastocysts .....................................79
Nuclear condensation, DNA damage, and apoptosis .............................................................80
DISCUSSION ............................................................................................................................86
x
Embryo development, sex ratio, and blastocyst cell number .................................................87
Nuclear condensation, apoptosis, and DNA-damage .............................................................89
CHAPTER THREE .......................................................................................................................93
INTRODUCTION ......................................................................................................................94
MATERIALS AND METHODS ...............................................................................................98
Experimental design ...............................................................................................................98
Chemicals ...............................................................................................................................98
Oocyte collection and in vitro oocyte maturation ..................................................................98
RNA extraction .......................................................................................................................98
Gene expression analysis .......................................................................................................99
Statistical analyses ................................................................................................................100
RESULTS.................................................................................................................................103
mRNA expression in GV and MII oocytes ..........................................................................103
Genes involved in oocyte maturation and spindle assembly ................................................103
Hormone receptor gene expression ......................................................................................104
Genes involved in stress and metabolism .............................................................................104
DISCUSSION ..........................................................................................................................111
mRNA expression in GV and MII oocytes ..........................................................................112
Genes involved in oocyte maturation and spindle assembly ................................................112
Hormone receptor .................................................................................................................114
Stress and metabolism genes ................................................................................................116
GENERAL DISCUSSION ..........................................................................................................118
SUMMARY AND CONCLUSIONS ..........................................................................................133
FUTURE DIRECTIONS .............................................................................................................135
REFERENCES ............................................................................................................................137
xi
LIST OF TABLES
Table 1. qPCR primers for sexing analysis ....................................................................................75
Table 2. Effect of IVM treatments on development, sex ratio, and total cell number of bovine
embryos ...................................................................................................................................82
Table 3 List of primers used for qPCR ........................................................................................101
xii
LIST OF FIGURES
Figure 1. Mean BPA concentrations of oocytes following IVM in their respective
treatment groups. Concentration of BPA (mean ± SEM) in oocytes at 24 hours following
oocyte maturation in IVM media supplemented with 0.1% ethanol (n=150), 15 ng/mL BPA
(n=150), or 30 ng/mL BPA (n=150). Analysis of variance (ANOVA) and Tukey’s multiple
comparison test, **p<0.01. .....................................................................................................52
Figure 2. Mean BPA concentrations of IVM media before and after incubation with
oocytes. Concentration of BPA (mean ± SEM) in IVM media with no supplementation
(IVM), or supplemented with 0.1% ethanol, 15 ng/mL BPA, or 30 ng/mL BPA prior to (T0)
or at 24 hours following oocyte maturation (T24+). Oocytes were removed prior to
processing of T24+ media. White asterisk = differences between T0 media samples of
different treatment. Black asterisks = differences between T0 and T24+ media samples of
the same treatment. (Analysis of variance (ANOVA) and Tukey’s multiple comparison test,
*p<0.05). .................................................................................................................................53
Figure 3. Representative images of meiotic stages during bovine oocyte maturation.
Representative images of bovine oocytes during maturation with α-tubulin in green and
chromatin in blue. Stages include (A) germinal vesicle; (B) germinal vesicle breakdown; (C)
prometaphase I; (D) metaphase I; (E) anaphase I; (F) telophase I; (G) prometaphase II; (H)
metaphase II with polar body (arrow) visible. Scale bar: 20 = μm ........................................54
Figure 4. Proportion of oocytes to reach MII following IVM in their respective treatment groups.
Proportion of oocytes (mean ± SEM) to reach the MII stage following 24 hour incubation in
IVM media without (IVM; n=98) or with 0.1% ethanol (n=97), 15 ng/mL BPA (n=110), or
30 ng/mL BPA (n=108) supplementation. Fisher’s exact, *p<0.05. ......................................55
Figure 5. Representative classifications of MII oocytes for spindle morphology and chromosome
alignment. Representative images of bovine MII oocytes with α-tubulin in green (left
panels), chromatin in blue (middle panels) and merged images (right panels). Oocytes with
normal spindle morphology and chromosomes alignment (A), abnormal spindle morphology
(B), and chromosome dispersal (C) are displayed. Rows B and C represent spindle
abnormalities observed in oocytes treated with 30 ng/mL BPA. Visible polar body is marked
with an arrow. Scale bar = 20 μm. ..........................................................................................57
Figure 6. Proportion of MII oocytes displaying normal and abnormal spindle morphology
following IVM in their respective treatment groups. Proportion of oocytes in each of the
treatment groups displaying abnormal spindle morphology following 24 hour incubation in
IVM media without (IVM; n=25) or with 0.1% ethanol (n=21), 15 ng/mL BPA (n=32), or 30
ng/mL BPA (n=28) supplementation. Fisher’s exact, *p<0.05. .............................................58
Figure 7. Proportion of MII oocytes displaying aligned and dispersed chromosomes at the
metaphase plate following IVM in their respective treatment groups. Proportion of oocytes
xiii
in each of the treatment groups displaying dispersed chromosomes at the metaphase plate
following 24 hour incubation in IVM media without (IVM; n=26) or with 0.1% ethanol
(n=21), 15 ng/mL BPA (n=33), or 30 ng/mL BPA (n=31) supplementation. Fisher’s exact,
**p<0.01. ................................................................................................................................59
Figure 8. Representative images of test samples (S), and positive (+) and negative (-) controls.
Scale bar = 50 μM ...................................................................................................................77
Figure 9. Condensed nuclei (white arrows) in blastocysts. Representative images (A) and mean
proportions (B) of condensed nuclei without TUNEL staining in blastocysts in each of the
treatment groups. Mean ± SEM. Scale bars = 50 μM.............................................................83
Figure 10. DNA-damaged nuclei (white arrow) in blastocysts. Representative images (A) and
mean proportions (B) of TUNEL-positive, non-condensed nuclei in blastocysts in each of
the treatment groups. Mean ± SEM. Analysis of variance (ANOVA) and Tukey’s multiple
comparison test, *p<0.05. Scale bars = 50 μM .......................................................................84
Figure 11. Apoptotic nuclei (white arrows) in blastocysts. Representative images (A) and mean
proportions (B) of TUNEL-positive, condensed nuclei in blastocysts in each of the treatment
groups. Mean ± SEM. Analysis of variance (ANOVA) and Tukey’s multiple comparison
test, **p<0.01. Scale bars = 50 μM ........................................................................................85
Figure 12. mRNA expression profiles of GV and MII oocytes in ERβ (A), GLUT1 (B), HSP70
(C), CDC2 (D), TRβ (E), TUBA (F), DAZL (G), p53 (H), KIF5b (I), and AURKA (J).
Unpaired t-test; (*) indicates statistical significance, *p<0.05; **p<0.01. Experiments were
conducted on three different pools of 40 oocytes (n = 3) and replicated three times (r = 3) for
each sample ...........................................................................................................................105
Figure 13. mRNA expression levels (Mean ± SEM) of CDC2 (A), AURKA (B), KIF5b (C),
DAZL (D), and TUBA (E) in MII oocytes following IVM in respective treatment groups.
One-way ANOVA and Tukey’s multiple comparison test. Bars with different letters indicate
statistical significance (p<0.05; **p<0.01). Experiments were conducted on three different
pools of 40 oocytes (n = 3) and replicated three times (r = 3) for each sample....................106
Figure 14. mRNA expression levels of ERβ (A) and TRβ (B) in MII oocytes following IVM in
respective treatment groups. One-way ANOVA and Tukey’s multiple comparison test. Bars
with different letters indicate statistical significance (p<0.05). Experiments were conducted
on three different pools of 40 oocytes (n = 3) and replicated three times (r = 3) for each
sample. ..................................................................................................................................107
Figure 15. mRNA expression levels of ERβ (A) and TRβ (B) in blastocysts arising from oocytes
matured in vitro in respective treatment groups. One-way ANOVA and Tukey’s multiple
comparison test. Bars with different letters indicate statistical significance (p<0.05).
Experiments were conducted on three different pools of 40 oocytes (n = 3) and replicated
three times (r = 3) for each sample .......................................................................................108
xiv
Figure 16. mRNA expression levels of HSP70 (A), p53 (B), and GLUT1 (C) in MII oocytes
following IVM in respective treatment groups. One-way ANOVA and Tukey’s multiple
comparison test. Bars with different letters indicate statistical significance (p<0.05).
Experiments were conducted on three different pools of 40 oocytes (n = 3) and replicated
three times (r = 3) for each sample .......................................................................................109
Figure 17. mRNA expression profiles of HSP70 (A), p53 (B), and GLUT1 (C) in blastocysts
arising from oocytes matured in vitro in respective treatment groups. One-way ANOVA and
Tukey’s multiple comparison test. Bars with different letters indicate statistical significance
(p<0.05). Experiments were conducted on three different pools of 40 oocytes (n = 3) and
replicated three times (r = 3) for each sample ......................................................................110
xv
LIST OF ABBREVIATIONS
AI – anaphase I
ART – assisted reproductive technology
AURKA – aurora kinase A
BPA – bisphenol A
CDK1 – cyclin-dependent kinase 1
CDC2 – see CDK1
COC – cumulus oocyte complex
DAZL – deleted in azoospermia-like
DES – diethylstilbestrol
EDC – endocrine-disrupting chemical
EGA – embryonic genome activation
ERα – estrogen receptor alpha
ERβ – estrogen receptor beta
FSH – follicle stimulating hormone
GLUT1 – glucose transporter 1
GV – germinal vesicle
GVBD – germinal vesicle breakdown
hCG – human chorionic gonadotropin
HPG – hypothalamus-pituitary-gonad axis
HSP – heat shock protein
ICM – inner cell mass
IVC – in vitro culture
xvi
IVF – in vitro fertilization
IVM – in vitro maturation
IVP – in vitro produced
KIF5b – kinesin family 5b
LH – luteinizing hormone
LOAEL – lowest observed adverse effect level
MBT – mid-blastula transition
MET – maternal to embryonic transition
MI – metaphase I
MII – metaphase II
MTOCs – microtubule organizing centers
MZT – maternal-zygotic transition
N-CoR – nuclear receptor corepressor
OP – octylphenol
PMII – prometaphase II
ProI – prometaphase I
ROS - reactive oxygen species
T3 – triiodothyronine
T4 - thyroxine
TCDD – 2,3,7,8-tetrachlorodibenzodioxin
TE – trophectoderm
THs – thyroid hormones
TI - telophase I
xvii
TRα – thyroid receptor alpha
TRβ – thyroid receptor beta
TUBA – alpha tubulin
VEGF – vascular endothelial growth factor (VEGF)
INTRODUCTION
The oocyte and early embryo are particularly sensitive to their immediate surroundings.
The composition of follicular fluid greatly influences the developmental potential of oocytes and
resulting embryos, and alterations may result in decreased fertility of the mother, or have longterm, permanent effects on the offspring (Eppig et al., 1996; Carabatsos et al., 2000; Garrido et al.,
2000; Voronina & Wessel, 2003; Da Broi et al., 2013). Alterations in the oocyte’s
microenvironment (follicular fluid or in vitro maturation media) can influence oocyte metabolism
and developmental competence, gene expression patterns, and blastocyst quality and implantation
as well as pregnancy success (Leroy et al., 2012). Furthermore, evidence suggests that the
environment of the oocyte may effect adult health of the resulting offspring, as well as the
subsequent generation (Wolstenholme et al., 2012; reviewed by Krisher, 2013). The components
of follicular fluid impact oocyte maturation and embryonic development since oocytes rely on
external signals from their environment for growth, maturation, fertilization capability, and
developmental potential (Zheng et al., 2003; Aardema et al., 2013; Krisher, 2013). Thus the
ovarian follicle represents a fragile microenvironment which strongly influences oocyte
competence.
Variables in follicular fluid such as hormone levels, temperature, free radicals, fatty acid
levels, and glucose levels can alter oocyte maturation and alter embryonic developmental potential
(Kreiner et al., 1987; Hodges et al., 2002; Leroy et al., 2006; Marei et al., 2010; Aardema et al
2013; Valckx et al., 2014). Alterations of follicular fluid can also have negative effects on oocyte
maturation and early embryo development by activating stress pathways such as is observed in
heat stress, oxidative stress, and metabolic stress (Hodges et al., 2002; Robker et al., 2009; Leroy
et al., 2012). Since oocyte quality appears to be easily perturbed, a growing concern is how
1
environmental exposure to chemicals that have been detected in follicular fluid, may affect fertility
and early development.
Endocrine disrupting chemicals (EDCs) are a group of chemicals which can disrupt normal
endocrine functions by mimicking or antagonizing the actions of hormones (Colborn et al., 1993).
EDCs may alter fertility through disruption of the hypothalamus-pituitary-gonad (HPG) feedback
loop, or by their presence in the follicular fluid where they can act directly on the oocyte and
surrounding cumulus cells (Craig et al., 2011; Fowler et al., 2012; Qin et al., 2013). In general,
EDC exposure to oocytes during maturation results in decreased fertilization rate and decreased
developmental potential (Petro et al., 2012). Interestingly, some EDCs such as octylphenol (OP)
appear to be acting through alternative mechanisms than direct estrogen disruption (Pocar et al.,
2003). In recent years it has been recognized that EDCs may alter fertility by acting as
environmental stressors (Wang et al., 2012; Krisher, 2013). Thus EDCs may affect oocyte
developmental competence through hormonal pathways, alternative pathways such as stress
activation, or via multiple mechanisms. This is why it has been difficult to fully elucidate the
mechanism of EDCs, such as bisphenol A (BPA), in early development, and why controversy
remains in the field.
BPA was first synthesized in 1891 and its estrogenic properties first discovered in the
1930s (Dodds & Lawson, 1936; reviewed by Wolstenholme et al., 2011). BPA would have been
used as a pharmaceutical if it were not for the discovery that diethylstilbestrol (DES) was much
more potent than BPA (Dodds & Lawson, 1936). BPA’s other properties made it central in the
development of polycarbonate plastics and epoxy resins (Ortiz et al., 2009). As such, BPA is now
a common component of many consumer products that we come into contact with every day
(Vandenberg et al., 2007; Ye et al., 2009). It is not surprising therefore that in one study in 2005
2
BPA was detected in >95% of an American reference population (Calafat et al., 2005) and >97%
detection was recorded in 2011 (Braun et al., 2011). BPA levels have also been detected in
follicular fluid of women at an average level of 2.4 ng/mL (Ikezuki et al., 2002). In vitro and in
vivo studies have demonstrated BPA exposure to the cumulus oocyte complex (COC) can result
in oocyte abnormalities such as aneuploidy and meiotic arrest (Hunt et al., 2003; Can et al., 2005;
Susiarjo et al., 2007; Lenie et al., 2008; Machtinger et al., 2013). However, the mechanism by
which BPA exposure to oocytes during maturation affects the earliest life stages is not as clear.
Early developmental changes that occur as a result of environmental exposures may give an idea
of how these chemicals exert their effects.
Oocyte quality and developmental competence are critical in the establishment of a healthy
pregnancy (Leroy et al., 2012). Examination of effects at the oocyte and early embryo stages shed
light on how maternal exposure to BPA may impact fertility and offspring health. The goal of this
thesis is to evaluate the effects of BPA exposure during bovine oocyte maturation in vitro on
aspects relating to developmental competency and quality of the oocyte and early embryo. Nuclear
maturation of the oocyte, and the integrity of the metaphase II spindle were assessed to determine
the effects of BPA on oocyte quality and embryo developmental potential. Secondly, the
preimplantation embryo was assessed following oocyte maturation exposures to evaluate
embryonic development, by calculating development rates, and embryo quality by evaluating
apoptosis, sex ratio, and total cell number of blastocysts. Finally, gene expression was evaluated
in both oocytes and blastocysts to further elucidate the actions of BPA and how early
developmental changes may be regulated at the transcriptional level.
3
LITERATURE REVIEW
Mammalian oogenesis
Oocytes provide both the maternal portion of genetic material, as well as cytoplasmic
components necessary for development of the oocyte itself and the early embryo. The oocyte pool
is established during gestation, and the life-time supply of oocytes is present in the ovaries at birth
contained within ovarian primary follicles. The follicle houses the oocyte throughout its
development, and the follicular environment in which the oocyte grows and matures is critical to
developmental success. Oocyte development involves highly complex processes consisting of
signalling between the oocyte and somatic cells of the follicle. These interactions allow for the
regulation of developmental processes such as oocyte metabolism, fertilization, and cell cycle
progression (reviewed by Li & Albertini, 2013). These events, brought on by cell-cell
communication, particularly that between oocytes and granulosa cells, are essential for the
initiation and maintenance of early embryo development (reviewed by Li & Albertini, 2013).
Primary oocytes contained within the follicles that are present from birth are arrested at
prophase I, and are maintained in this state by inhibitory signals from cumulus cells surrounding
the oocyte (Norris et al., 2009). The primary follicles remain dormant until the onset of menarche,
the first egg release. Continuation of development is signalled by somatic cells of the follicle
(reviewed by Voronina & Wessel, 2003). The oocyte begins to grow within the follicle, but
remains meiotically arrested, and is characterized by an enlarged nucleus, the germinal vesicle
(GV). During the growth phase, the oocyte accumulates and stores large amounts of maternal
mRNAs, proteins, and organelles that are invaluable for the growth and maturation of the oocyte
(Hunter & Moor, 1987; Wessel et al., 2001; Krisher, 2013). Following fertilization, these stores
function to support and regulate early embryonic development, particularly until the maternal to
4
embryonic transition (MET), where embryonic transcription takes over (Voronina & Wessel,
2003; Krisher, 2013). Thus the maternal mRNAs and proteins accumulated during the growth
phase of oogenesis are critical factors in oocyte competence (Paczkowski et al., 2011; Krisher,
2013). Following the growth phase, the oocyte will begin maturation, another important period of
time in the oocyte’s development.
Oocyte maturation
The LH surge which leads to ovulation initiates oocyte maturation which is, in part,
characterized by resumption of meiosis from the germinal vesicle stage. Oocyte maturation is a
critical period of oocyte development that dictates the developmental potential of the oocyte. It is
a highly coordinated process comprising of a series of changes in the oocyte’s nucleus and
cytoplasm. These changes are referred to as nuclear and cytoplasmic maturation. Both are essential
to oocyte quality as these events directly influence the success of fertilization and embryo
development (Combelles & Albertini, 2002).
Nuclear maturation is the resumption of meiosis from prophase I to metaphase II (MII).
The oocyte is arrested at this phase until fertilization occurs, or it is degraded without having been
fertilized. Nuclear maturation includes germinal vesicle breakdown (GVBD), chromosome
condensation, assembly of the meiotic spindle, and ultimately the formation of a large secondary
oocyte as well as the first polar body. The metaphase I (MI) spindle is positioned cortically so that
loss of cytoplasm during the extrusion of the first polar body is minimal (Brunet & Verlhac, 2010).
Following extrusion of the first polar body, the chromosomes which remain in the oocyte continue
to progress through meiosis until their arrangement on the meiotic spindle at MII (reviewed by
Voronina & Wessel, 2003). Nuclear maturation is complete at this stage, however meiosis II is not
5
complete until fertilization and extrusion of the second polar body has occurred (reviewed by Li
& Albertini, 2013).
The resumption of meiosis and the completion of nuclear maturation are linked to
developmental competence. Decreased maturation success, or incomplete meiosis progression are
indicators of poor quality oocytes. An oocyte unable to resume and complete maturation under
proper conditions is of poor quality and has not obtained the necessary factors during growth that
allows it to initiate and sustain maturation. Suboptimal conditions can also lead to an oocyte’s
inability to resume and complete maturation, and has been associated with a decrease in the
proportion of oocytes reaching nuclear maturity. Thus the ability of an oocyte to complete nuclear
maturation may be used as an indicator of oocyte quality and/or the quality of environment in
which the oocyte is maturing. Oocytes of good competence may be able to withstand
environmental perturbations, whereas lower quality oocytes may not be able to survive under such
conditions. Furthermore, the competence of oocytes that are able to reach maturity may be
compromised, possibly resulting in long-term detrimental effects on the embryo and offspring
health. An important marker of oocyte competence of mature oocytes is the MII spindle.
The MI and MII spindles are critical to the completion of meiosis. The spindle is composed
of bundles of microtubules that attach to kinetochores of chromatin, and guide the chromosomes
during segregation. At MII, the spindle is barrel- or ovoid-shaped with the microtubules extending
inwards from two microtubule organizing centers (MTOCs) to the chromosomes aligned
equatorially along the metaphase plate. Proper positioning of the spindles helps to minimize the
loss of ooplasm during polar body extrusion (Brunet & Verlhac, 2010), and the risk of aneuploidy
that results from improper segregation of the chromosomes (Volarcik et al., 1998). Human oocytes
have been found to exhibit a relatively high incidence of errors in chromosome segregation, and
6
these errors have been linked to pregnancy failure, embryonic abnormalities as well as genetic
diseases (Hassold & Hunt, 2001; Li & Albertini, 2013).
Nuclear maturation is one of two important components that together comprise oocyte
maturation. Cytoplasmic maturation, though less well defined, plays a critical role in preparation
of the cell to support fertilization and early embryonic development. Cytoplasmic maturation
consists of various cytoplasmic rearrangements. These changes allow the oocyte to support
fertilization and commence embryonic development (reviewed by Krisher, 2013). Some important
events that occur during cytoplasmic maturation include metabolism of carbohydrates and lipids,
rearrangement of the mitochondria, oxygen radical reduction, epigenetic programming, growth
factor secretion, and cross-talk between the oocyte and its surrounding cumulus cells (reviewed by
Krisher, 2013). Cytoplasmic maturation is thus also an important determining factor for oocyte
quality and ideally should be considered when assessing oocyte quality as well as nuclear
maturation. However the only current method of evaluating the completion of cytoplasmic
maturation is successful fertilization and positive developmental outcome of embryonic
development. Thus nuclear maturation is largely assessed in efforts to determine if oocyte
maturation has been successful.
Oocyte maturation is a particularly critical phase of the oocyte’s development and the
establishment of developmental competency. The environment in which the oocyte matures can
have significant and long-term impacts on the resulting embryo’s ability to sustain development
(Combelles & Albertini, 2002), as well as the health of resulting offspring. Because of the highly
complex signalling that occurs in the follicular environment, any alterations made to that
environment that may disrupt proper signalling can have detrimental effects on the maturing
oocyte.
7
Oocyte competence
Prior to fertilization and embryonic development, the oocyte must acquire developmental
competency, defined by various cellular and molecular properties, that enable the oocyte to resume
and complete meiosis, support fertilization and oocyte activation, initiate mitosis following
fertilization, and support embryonic development until the time at which embryonic transcripts
take over (First et al., 1988; Albertini et al., 2003; Krisher, 2013). The process through which these
competencies are acquired is lengthy and complex, with important events affecting competency
occurring late in the growth phase and during oocyte maturation.
Oocyte competence, or quality, is critical to female fertility, having both short- and longterm implications regarding embryonic survival and implantation, fetal development, and adult
health of the offspring. The oocyte is particularly vulnerable and oocyte competence can be
affected both by internal and external factors. Internal factors such as gene transcription, protein
translation, metabolism, and organization of the cytoplasm contribute to the competence of the
oocyte, and competence can be further influenced by the external environment to which the oocyte
is exposed.
The importance of spindle function with respect to oocyte quality has long been accepted.
The MII spindle is therefore often studied as an indicator of oocyte quality due to its role in meiosis
completion, polar body extrusion, and segregation of the chromosomes. A bipolar spindle with
normal microtubule assembly is necessary for proper chromosome segregation during meiosis,
with improper chromosome segregation at this time resulting in aneuploidy which can cause
chromosomal disorders such as Down’s syndrome, or errors that cannot be sustained and lead to
pregnancy loss (Hassold & Hunt, 2001; Mtango et al., 2008). Chromosomal aneuploidies are
common, and the proportion of aneuploid cells within an embryo directly correlates with the
8
developmental potential of the embryo (Baltaci et al., 2006; Mtango et al., 2008).
In human oocytes, flattening of both spindle poles was associated with increased incidence
of misalignment of the chromosomes at the metaphase plate (Coticchio et al., 2013). An
association has been found in mammalian models between spindle abnormalities and aneuploidy
incidence (Sanfins et al., 2003; Bromfield et al., 2009). For instance, abnormalities in spindle
morphology and chromosome alignment at the metaphase plate have been found to be predictors
of aneuploidy in aging women (Battaglia et al., 1996). In this study, the spindle of oocytes from
older women, in comparison to those from younger women, exhibited a significantly higher
incidence of abnormal tubulin placement within the spindle as well as chromosomal displacement
from the metaphase plate at MII. Battaglia et al. (1996) speculated that this increase in spindle
abnormalities leads to the high prevalence of aneuploidy in oocytes of older women.
The follicular environment
The mammalian oocyte grows and matures within a follicle, and the environment of that
follicle, which is reflective of the external maternal environment, is extremely important in the
developmental fate of the oocyte. The follicle provides nutrients necessary for the growth of the
oocyte, and produces signals that regulate oocyte development (Eppig et al., 1996; Carabatsos et
al., 2000; Voronina & Wessel, 2003). Follicular growth, as well as oocyte competence, are
controlled by complex signalling and communication between theca, granulosa, and cumulus cells
within the follicle. Bidirectional signalling in the form of autocrine, paracrine, and endocrine
regulation as well as gap junctional interactions, are responsible for regulating growth and
development of the oocyte and the follicular cells (Voronina & Wessel, 2003). The oocyte plays a
dominant role, secreting factors that regulate follicular development and the function of cumulus
9
cells (Eppig et al., 2002; Matzuk et al., 2002). Conversely, the cumulus cells help to ensure proper
growth and maturation of the oocyte by transporting essential molecules to the oocyte (Eppig,
2001; Gilchrist et al., 2008). Therefore, oocyte competence is acquired both by the oocyte’s ability
to produce and secrete the necessary factors, as well as the response of the cumulus cells.
Follicular fluid is vital to the success of oocyte maturation and alterations to the
composition of follicular fluid have the potential to affect oocyte maturation and the quality of
oocytes, possibly by interfering with these complex signalling interactions. For instance, Da Broi
et al. (2013) found that follicular fluid from infertile women with endometriosis, which has been
reported to differ in comparison to follicular fluid from fertile women (Garrido et al., 2000),
resulted in increased incidence of meiotic abnormalities.
Hormone levels in follicular fluid play an important role in the developmental competency
of the oocyte. The levels of E2 (Aardema et al., 2013), progesterone (Aardema et al., 2013), and
growth hormone (Modina et al., 2007) can be predictors of blastocyst production and markers of
oocyte competence. E2 and progesterone levels in preovulatory follicles were found to be
predictors of the developmental potential of oocytes in heifers (Kreiner et al., 1987; Aardema et
al., 2013), and were also found to improve developmental competence when added to in vitro
maturation (IVM) culture media (Zheng et al., 2003). Furthermore, follicular fluid concentrations
of growth hormone (Modina et al., 2007), FSH (Suchanek et al., 1988), hCG (Ellsworth et al.,
1984), and LH (Cha et al., 1986) have all been associated with oocyte competence, with levels of
FSH, hCG, and LH being positively correlated with oocyte competence in humans, and levels of
growth hormone inversely related to oocyte competence in cows.
The complex milieu of factors contained within the follicular environment explains the
difficulty assisted reproductive technologies (ARTs) have had in creating successful pregnancies
10
from oocytes that have been matured in vitro (reviewed by Chian, 2004). In domestic animals, on
the other hand, in vitro oocyte maturation is more commonly used and has resulted in greater
success in the production of offspring from embryos which were derived from in vitro matured
oocytes (Rodriguez-Martinez, 2012). However, pregnancy rates from in vitro matured oocytes
remain substantially lower than oocytes which matured in vivo (Rizos et al., 2002). Furthermore,
environmental conditions of the oocyte, in vivo and in vitro, can have significant impacts on oocyte
quality and developmental potential (reviewed by Krisher, 2013). Thus viability of an oocyte and
resulting embryo can be strongly influenced by the environment in which the oocyte matures.
Environmental influences
The follicular environment is reflective of the external environment experienced by the
female and the maternal environment can have a direct impact on oocyte competence.
Environmental factors that can lead to a decrease in developmental competency include oxidative
stress, heat stress, maternal hormone status (Hodges et al., 2002), in vitro culture conditions
(Trounson et al., 2001) and interruptions to normal oocyte-somatic cell signalling (Carabatsos et
al., 1998). Of particular relevance to the current work are EDCs.
EDCs are environmental chemicals that are commonly encountered and may impact female
fertility; most are man-made chemicals that interfere with endogenous hormone functioning.
Certain EDCs may alter endogenous hormone synthesis and secretion, metabolism, transport, and
ligand action. Most commonly, EDCs interfere with estrogenic pathways, exhibiting estrogen-like
actions by binding to and activating the estrogen receptors. EDCs can interfere with normal
reproductive functioning at various stages of the reproductive life cycle. Oocyte, embryonic and
fetal developmental period are of particular concern, with exposure during this time potentially
11
leading to significant reproductive detriments that may be passed on to the subsequent generation
(reviewed by Krisher, 2013). The severity of effects as a result of developmental exposure depends
on the nature of the specific EDC, the exposure level and frequency to the EDC, and the timing of
exposure during development. EDCs may affect oocyte competence in a number of ways. They
may act directly on the oocyte, or indirectly either via the follicular cells, or disrupting the
hypothalamic-pituitary-gonadal axis to interfere with the secretion of gonadotropins (reviewed by
Krisher, 2013).
The impact that the external environment has on the oocyte may also vary depending on
the inherent competence of the oocyte. For instance, oocytes with poor inherent competence may
be more severely impacted by poor environmental conditions than those oocytes with good
inherent competence. The oocyte must withstand environmental insults while retaining its
developmental competency in order to successfully perform its reproductive function. Poor quality
oocytes may be unable to properly support embryo development, and adverse developmental
outcomes that may be passed on to future generations are significantly affected by the quality of
the oocyte from which they developed (reviewed by Mtango et al., 2008). There are a number of
developmental implications of poor oocyte competence. Maturational effects include a decrease
in maturation success, altered gene expression of the cumulus oocyte complex (COC), disruption
of the meiotic spindle assembly, changes in distribution of mitochondria, chromosome dispersal
at the metaphase plate, and chromosomal abnormalities such as aneuploidy. Poor developmental
outcomes such as these can be used to assess the quality of the oocyte from which an embryo
developed.
12
Early embryo development
Following fertilization the ovum has become a single diploid cell, referred to as a zygote.
The zygote is enclosed within the zona pellucida, an acellular glycoprotein membrane surrounding
the embryo, and undergoes its first mitotic cleavage resulting in a 2-cell embryo (Hardy et al.,
1989). The early embryo undergoes rapid mitotic division cycles, during which the amount of
DNA is exponentially increasing while the amount of cytoplasm in the embryo remains constant
eventually becoming a morula of at least 16 cells. At the morula stage the cells are compacted
(Hardy et al., 1989), but remain undifferentiated. Cellular differentiation and cavitation of the
embryo occur to form the early blastocyst. Embryonic cells differentiate into trophoectoderm cells
(TE cells) and the inner cell mass (ICM) that form the blastocyst, and fluid accumulates in a
blastocoel cavity (Hardy et al., 1989). The early blastocyst in bovine is reached 7-8 days post
fertilization. The blastocyst, although small at first, begins to expand. Eventually the blastocyst
will hatch from the protective zona pellucida in preparation for implantation into the uterine
endometrium in humans, or attachment in bovine.
During the early cell cycles, the cells of the embryo are undifferentiated and
transcriptionally silent (reviewed by Vigneault et al., 2009) and maternally provided proteins and
mRNAs regulate development (reviewed by Schier, 2007). The embryonic genome becomes
transcriptionally active at later cell cycles during early embryo development. This period of time
is referred to as the MET, and has also been called the maternal-zygotic transition (MZT),
embryonic genome activation (EGA), and mid-blastula transition (MBT) (reviewed by Schier,
2007). The MET is a critical period of early embryo growth. The maternal products begin to
degrade, embryonic transcription is initiated and the embryo becomes dependent on expression of
embryonic genes (reviewed by Memili & First, 1999). Prior to the MET, the embryo is largely
13
dependent on maternal RNAs and proteins synthesized during oogenesis (reviewed by Memili &
First, 1999). The timing of the MET varies between species, but similarly it is a transitional period
at which time the transcriptional contribution provided by the mother begin to decline, and that of
the developing embryo begin to increase (reviewed by Memili & First, 1999). In bovine embryos,
genomic activation occurs at the 8- to 16-cell stage (reviewed by Vigneault et al., 2009).
RNAs stored during oogenesis are important in sustaining embryonic development before
embryonic transcription takes over (reviewed by Memili & First, 1999). This can thus be a
vulnerable period for the embryo if it is not well equipped to take control of its genomic activity.
This also sheds light on the importance of oocyte competence. The embryo must be supplied with
sufficient transcripts and proteins acquired during oocyte growth and maturation to survive until
the embryonic genome takes over. Furthermore, alterations in the gene and protein expression
profiles of mature oocytes may affect early embryonic development (Evsikov et al., 2006),
possibly resulting in embryonic arrest, cell death, and/or poor developmental outcome.
Vulnerability of the embryo
Successful development of an early embryo will largely depend on its environment as well
as its innate ability to respond to environmental changes or stressors. Early embryos are void of
defense systems such as immune or nervous systems that are present in the adult. Parental defenses
are in place to provide some protection to the embryo, however the embryo still remains vulnerable
to certain types of stresses in its environment. The embryo has developed its own defense
mechanisms which increase its chances of survival. In real world conditions, embryo development
is stable and early embryos are equipped with high levels of cellular defenses in preparation for
environmental changes (reviewed by Hamdoun & Epel, 2007). These cellular defenses are present
14
in the egg prior to fertilization. Later on in preimplantation development, the embryo may be able
to buffer stress to adapt to the environmental conditions (reviewed by Hamdoun & Epel, 2007).
Whether a particular maternal stress affects embryonic survival depends not only on the magnitude
of the stress imposed on the embryo, but also on the effectiveness of the embryonic adaptive
response to that stress.
In vivo embryo development is difficult to duplicate in vitro. Sub-optimal culture
environments are thought to be responsible for the high frequency of early developmental failure
that is associated with in vitro-produced embryos (Johnson & Nasr-Esfahani, 1994; Betts & King,
2001). Over recent years much research has gone into improving the in vitro fertilization (IVF)
system in hopes of improving cleavage, embryo developmental potential, blastocyst development,
and ultimately successful pregnancy and live birth. However it is not only in vitro systems that can
hinder early embryo development. Various conditions of stress in vivo can also result in
consequences to the embryo including delayed development, apoptosis induction, altered gene
expression, and miscarriage (Wilson et al., 1985; Arck, 2004; Dennery et al., 2007; Ornoy, 2007).
These stresses include hypoxia, changes in temperature, pathogens, UV radiation, free radicals,
environmental toxins, and maternal diet and health (reviewed by Hamdoun & Epel, 2007). The
developmental fate of the embryo rests upon the embryo itself and its defense capabilities of
withstanding such stressors.
The ability of embryonic developmental programs to withstand stressors that have the
potential to disrupt cellular and molecular mechanisms of embryogenesis dictates the severity of
the effects caused by the stressor (Hamdoun & Epel, 2007). These programs are evidenced by the
survival and maintenance of embryonic viability despite exposure to potential stressors (Hamdoun
& Epel, 2007). The evolution of embryonic defenses was shaped by environmental perturbations
15
expected to be encountered by the embryo, and resulted in embryos with stronger defense systems
against commonly encountered stressors (Hamdoun & Epel, 2007). Embryonic robustness,
however, can be overwhelmed if the stress exceeds the limit than can be buffered by the embryo
(Hamdoun & Epel, 2007). A growing concern is that man-made chemical stressors currently
encountered by embryos may be beyond the range of these protective mechanisms, and may lead
to poor developmental outcomes.
Apoptosis in the preimplantation embryo
Apoptosis is a form of cell death that is critical to normal development. Ineffective
apoptotic mechanisms can lead to disorders of the central nervous system, and webbed hands and
feet. Apoptosis, or programmed cell death, is a structured series of events that is triggered by the
cell and essentially causes it to self-destruct. In doing so, the damaged cell can be destroyed
without affecting neighbouring cells. Apoptosis eliminates abnormal or damaged cells as well as
cells with abnormal developmental potential (Hardy, 1999; Betts & King, 2001; Paula-Lopes &
Hansen, 2002a). Apoptotic cell death results in the removal of these cells without inducing
inflammation, thus preserving the cellular integrity of neighbouring cells (Betts & King, 2001;
Paula-Lopes & Hansen, 2002b).
In the blastocyst, cell death may be a way to regulate development, eliminating abnormal or
damaged cells (Betts & King; 2001; Hansen & Fear, 2011). By removing cells that are damaged,
abnormal, or have an inappropriate developmental potential, apoptosis acts as a quality control
mechanism and is an important factor in animal development (Jacobson et al., 1997; Betts & King,
2001). The ability of the embryo to induce apoptosis in times of stress may help it to survive
suboptimal conditions, however the extent to which apoptosis occurs in the blastocyst plays an
16
important role in the fate of that embryo (Betts & King, 2001). Where mild induction of apoptosis
might help the embryo survive by eliminating damaged cells, severe stress may cause extensive
apoptosis that could compromise development and survival of the embryo (Byrne et al., 1999;
Betts & King, 2001; Paula-Lopes & Hansen, 2002b; Hansen & Fear, 2011). Thus the successful
embryonic development is dependent on the embryo’s ability to protect itself and respond to
stressors (Hamdoun & Epel, 2007).
Adverse environmental conditions during preimplantation development can increase the
proportion of cells undergoing apoptosis in the early embryo (Paula-Lopes & Hansen, 2002b).
Stressors encountered by preimplantation embryos include heat stress, oxidative stress, pathogens,
and exposure to toxic or xenobiotic chemicals (Betts & King, 2001; Hamdoun & Epel, 2007).
These environmental stressors have been found to induce apoptosis in early embryos under certain
conditions and may compromise embryonic viability and survival depending on the severity of the
stress, and the time of exposure during development (Paula-Lopes & Hansen, 2002a; Hamdoun &
Epel, 2007).
Embryo sex ratio
Mendelian genetics assumes a 1:1 sex ratio with equal proportions of males and females
born. However, variation in sex ratio is common and deviations from the 1:1 expectation are often
recorded in humans and other mammals. The natural sex ratio is thought to be about 1.05, 105
males born for every 100 females, or about 52.5% males. Studies have found that sex ratio can
shift as a result of various factors including fertility, nutrition, food availability, hormone status of
either parent, the timing of insemination in relation to ovulation, the use of certain ART techniques
in both humans (Menezo et al., 1999; Kallen et al., 2005; Luna et al., 2007; Dean et al., 2010) and
17
non-human mammals (Iwata et al., 2008).
Maternal stress has been found to impact the sex ratio of offspring in some cases. In
rodents, females exposed to social stress produced fewer sons in mice (Krackow, 1997), hamsters
(Pratt & Lisk, 1989), and rats (Lane & Hyde, 1973; Moriya et al., 1978), and the factors that may
result in a decrease in sex ratio (fewer males) includes, but is not limited to, stressors such as
pollution (James, 1998) and confinement stress (Krackow, 1997). Fewer males also tend to be
conceived in humans under suboptimal conditions, including environmental disasters (Fukuda et
al., 1998), pollution (Mocarelli et al., 2000; Weisskopf et al., 2003), and mothers of advanced age
(Orvos et al., 2001). This trend indicates that male embryos are more fragile and less likely to
survive under stressful conditions than female embryos. These observations comply with the
Trivers & Willard (1973) hypothesis that postulates an excess of male offspring is only favoured
by natural selection when conditions, and likelihood to survive, is good, and mothers in poor
conditions benefit most by the production of daughters (Trivers & Willard, 1973). This may be a
result of differential survival rates of male and female embryos during the preimplantation
development.
In vitro, blastocyst sex ratio has been found in many instances to be altered by
environmental conditions such as hyperglycemia, oxidative stress, in vitro culture media
composition, and exposure to EDCs. Increased glucose levels in in vitro culture media appears to
favour male embryos and inhibit female embryo development (Gutiérrez-Adán et al., 2001a;
Larson et al., 2001), whereas paternal exposure to the dioxin TCDD resulted in skewed sex ratio
in favour of female embryos (Ishihara et al., 2007).
These differences in response to environmental conditions are speculated to be a result of
metabolic, genetic, and epigenetic differences between male and female embryos that have been
18
observed in vitro. During the preimplantation period, male and female embryos differ only in the
content of their sex chromosomes, and differences exhibited by embryos at this time must be a
result of transcriptional dimorphism (reviewed by Bermejo-Alvarez et al., 2011). Furthermore,
epigenetic differences resulting from the presence of one versus two X-chromosomes may be a
driving force in sex differences and how embryos respond to environmental conditions (reviewed
by Gutiérrez-Adán et al., 2006).
Expression of genes encoded by the sex chromosomes, which may also affect autosomal
gene expression, differs between male and female preimplantation embryos (reviewed by
Bermejo-Alvarez et al., 2011). Y-linked genes are only expressed in male embryos, and X-linked
genes are expressed doubly in females. X-chromosome inactivation (XCI) compensates for this
disparity ensuring X-linked genes (in most cases) are transcribed equally in male and female adult
tissues (reviewed by Bermejo-Alvarez et al., 2011). However, during early embryo development
XCI is incomplete, or genes are reactivated, and female embryos exhibit higher expression of many
X-linked genes (Kobayashi et al., 2006; Bermejo-Alvarez et al., 2010a). This phenomenon is
exhibited in mouse (Kobayashi et al., 2006), bovine (Gutiérrez-Adán et al., 2000), and human
embryos (Taylor et al., 2001).
These transcriptional alterations are responsible for variations between male and female
embryos such as metabolic differences, and dimorphic susceptibilities to suboptimal in vivo and
in vitro conditions (reviewed by Gutiérrez-Adán et al., 2006). Resulting alterations in molecular
pathways, such as the pentose-phosphate pathway (PPP) that regulates glucose metabolism, may
result in varying susceptibilities of male and female embryos. For instance, male embryos have
been reported to metabolize glucose at a higher rate than females (Tiffin et al., 1991) and appear
to benefit more than females under hyperglycemic conditions in vitro; however the opposite has
19
also been reported (Jimenez et al., 2003). Conversely, female embryos exhibit increased
expression of X-linked genes such as those related to energy metabolism, the regulation of oxygen
radicals, and apoptosis inhibition (Gutiérrez-Adán et al., 2000; Jimenez et al., 2003). Since various
stressors can lead to embryonic overproduction of reactive oxygen species (ROS), female embryos
may be better equipped to survive such a stress due to enhanced ability to buffer the amount of
cellular ROS (Perez-Crespo et al., 2005). Furthermore, male embryos have been shown to be more
sensitive to oxidative damage induced by heat stress, which may be due to the increased expression
of X-linked genes in females that result in an increased ability to buffer environmental stress
(Perez-Crespo et al., 2005).
Therefore, whether the embryo possesses one or two X chromosomes may underlie the
differences in the early embryo (Gutiérrez-Adán et al., 2006). This may explain observations that
male embryos may be more vulnerable to stressors than female embryos under certain
environmental conditions. Since male and female embryos evidently respond differently to stress,
a skew of the natural sex ratio may be indicative of environmental stress imposed on the embryo.
Sex ratios have therefore been widely used as an indicator for reproductive health and the
embryonic environment.
Blastocyst gene expression
A milestone of preimplantation development is formation of the blastocyst. As discussed
earlier, the MET is a period of embryonic genome activation where transcripts from the mother
begin to degrade and those originating from the embryo increase and take over development.
During this period, the embryonic gene expression program is developed (reviewed by Schultz,
2005). Although early embryo development is autonomous, environmental conditions of the
embryo or the oocyte from which it developed can influence preimplantation development and, if
20
conditions are extreme, lead to abnormal embryonic development (Niemann & Wrenzycki, 2000).
Embryonic gene expression can be influenced by environmental factors including in vitro culture
conditions (Lonergan et al., 2006). Though not as thoroughly studied, the environment in which
the oocyte matures has also been found to alter transcript abundance in the mature oocyte
(Lonergan et al., 2003a) as well as in the blastocyst from which it develops (Russell et al., 2006).
Changes in mRNA transcript abundance at the blastocyst stage caused by environmental
perturbations may indicate an altered quality, or developmental competency of the blastocyst,
however the functional significance of variations in transcript abundance can be difficult to
interpret (reviewed by Duranthon et al., 2008). El-Sayed et al. (2006) analyzed differences in
transcript abundance of genes from embryos resulting in pregnancy versus those that did not.
Transcript abundance for genes involved in carbohydrate metabolism, implantation, and placental
development, among others, were found to be higher in embryos associated with successful
pregnancies whereas those of inflammatory cytokines, transcription factors, glucose metabolism,
and implantation inhibition were lower (El-Sayed et al., 2006).
It is evident that alterations to the in vitro culture environment of embryos can influence
the gene expression of a variety of genes in the blastocyst, however the influence of oocyte stress
on resulting blastocyst gene expression is controversial. Studies examining oocyte exposure tend
to focus on acute effects in the oocyte whereas blastocyst effects studied are normally a result of
environmental exposures during embryo development. Evidence has suggested that the
preimplantation period is the most important factor when considering blastocyst quality (reviewed
by Lonergan et al., 2006), however the environment of the oocyte has the capability of inducing
long term effects including changes in gene expression in the blastocyst (Pocar et al., 2001; Russel
et al., 2006). For instance, oocyte exposure during maturation to elevated non-esterified fatty acid
21
concentrations resulted in changes to embryonic gene expression and phenotype, possibly as a
result of altered metabolic strategies (Van Hoeck et al., 2013). Understanding the importance of
the environment in which the oocyte matures and embryo develops is critical in formulating a
comprehensive picture of how environment affects early development. Analyzing gene expression
changes in the blastocyst as a result of environmental perturbations provides an indication of the
embryo quality and stress that has been experienced by the embryo.
Hormone receptors
Nuclear hormone receptors are transcription factors that are activated when a ligand, such
as steroid or thyroid hormones (THs), binds to the receptor forming a complex that will bind to
DNA response elements contained within promoter regions of the target genes (Kinyamu &
Archer, 2003; reviewed by Rastinejad et al., 2013), and thereby regulate the gene expression of
their target genes (Mangelsdorf et al., 1995; Gronemeyer et al., 2004). Nuclear receptors play an
important role in the oocyte and early embryo (Beker-van Woudenberg et al., 2004), and are
essential for implantation/attachment (reviewed by Vasquez & DeMayo, 2013). Alterations in
hormone receptor mRNA may indicate poor developmental programming or improper conditions,
and may be predictive of undesirable reproductive outcomes.
E2, a steroid hormone, plays a critical role in fertility, oocyte development, and early
embryo development. The hormone receptors to which E2 binds in order to exert its effects, ERα
and ERβ, and the levels at which they are expressed can therefore impact the proficiency of these
programs and the viability of oocytes and embryos. ERα is expressed in cumulus cells whereas
ERβ is expressed both in cumulus cells and oocytes (Beker-van Woudenberg et al., 2004). In the
porcine embryo, ER mRNA is present in the early cleavage divisions, but is undetectable around
22
the 5- to 8- cell stages, and then reappears at the blastocyst stage (Ying et al., 2000; Chingwen et
al., 2000). In mice embryos, a similar pattern is observed with expression disappearing at the 8cell stage and reappearing at the morula and blastocyst stages (Hou & Gorski, 1993; Hiroi et al.,
1999). It has been suggested that the reappearance of ER mRNA at the blastocyst stage indicates
that E2 at this time may act directly on the embryo (Chingwen et al., 2000); this correlates with
the essential role of the hormone in the establishment and maintenance of pregnancy. Changes in
ER mRNA levels at blastocyst may therefore lead to reproductive deficits such as pregnancy loss.
Disruption of ERα expression in blastocysts did not affect embryo development and implantation
in mice (Saito et al., 2014), however Hou et al. (1999) suggested that the lack of natural ER
mutations may be evidence that ERs are essential for embryonic survival.
Thyroid hormones, T3 and T4, are required for normal growth and differentiation of most
organs in vertebrates (Darras et al., 2011). T3 and T4 regulate metabolism, cardiac function,
remodelling of bone and brain development (reviewed by Boas et al., 2012). However, the benefits
of THs precede organ development. THs have been found to be present in the female reproductive
tract and follicular fluid (Ashkar et al., 2010a), and beneficial to early embryo quality (Ashkar,
2013). THs exert their action by binding to the TH receptors (TRs) TRα and TRβ (reviewed by
Darras et al., 2011). TRs are expressed from the 2-cell to the blastocyst stage in bovine embryos,
suggesting active transcription of TR mRNA throughout preimplantation development (Ashkar,
2013). TH supplementation to in vitro culture (IVC) media did not alter TR mRNA expression of
the blastocyst, however TR expression may be altered by other exposures. For instance, TRβ
expression was suppressed as a result of bisphenol A (BPA) exposure in Xenopus embryos
(Iwamuro et al., 2003). As will be discussed later, BPA is a chemical with demonstrated antithyroid properties, and given the importance of THs during preimplantation development, more
23
research is needed regarding the disruption of TR expression during early development.
Embryonic stress, metabolism, and gene expression
As discussed earlier, stress incurred by the embryo can lead to increased incidence of
apoptosis and a skewed sex ratio. There are a number of genes in preimplantation development
that are involved in the embryonic response to stress. Two well-characterized examples of
embryonic stress include heat and oxidative stress, both of which have been shown to result in a
decrease in oocyte competence and embryonic developmental potential in mammals (Al-Katanani
et al., 2002; Sartori et al., 2002; Tamura et al., 2008; reviewed by Takahashi, 2012). Genes known
to be involved in the embryonic response to these stressors include the heat shock protein, HSP70,
the tumour suppressor, p53, and the glucose transporter, GLUT1. Transcription levels of these
genes have been shown to vary as a result of embryonic stress, though these responses appear to
be dependent on both the level of stress experienced and the stage of development at which the
stress occurs.
Heat shock proteins (HSPs) play an important role in oocyte fertilization and early embryo
development in mammals (Anderson, 1998; Neuer et al., 1998, 1999). The two major roles of
HSPs are that of molecular chaperones, and in the protection against cellular damage caused by
stress (Welch, 1992). HSP70 is a major heat shock protein that protects cells against detrimental
effects of stress (Welch, 1984; Hendrey & Kola, 1991) by preventing the denaturation of proteins
and inhibiting apoptosis (Matwee et al., 2001; Kregel, 2002). HSP70 plays a vital role in early
embryo development as evidenced by observations that exposure to HSP70 antibodies resulted in
decreased murine and bovine embryo development, and increased DNA fragmentation and
apoptosis (Neuer et al., 1998, 1999; Matwee et al., 2001). These results along with avian studies
24
which have demonstrated a correlation between developmental HSP70 expression and apoptosis
resistance during stress indicate that HSP70 plays an important role in apoptosis inhibition during
early embryo development (Bloom et al., 1998).
HSP70 transcription is increased in immature oocytes and embryos of Bos taurus as a result
of heat stress (in vivo) (Camargo et al., 2007) or heat shock (in vitro), respectively (Kawarsky &
King, 2001). Additionally, other stressors such as inadequate culture conditions of in vitro
produced (IVP) bovine embryos (Wrenzycki et al., 1999), and exposure of zebrafish (Danio rerio)
embryos to heavy metals and pesticides (Scheil et al., 2010) resulted in increased HSP70 mRNA
and protein expression, respectively. This protective capacity of HSP70, however, appears to be
developmentally acquired with embryos of at least the 8-cell stage exhibiting protective increases
in HSP70 expression (Kawarsky & King, 2001). HSP70 expression, though vital for normal
embryo development, appears to become increasingly critical in the face of adverse environmental
conditions experienced by the embryo. Expression of HSP70 mRNA is therefore often used as a
marker of stress in preimplantation embryos (Wrenzycki et al., 2001).
HSP70 interacts with the tumour suppressor protein p53, which is recognized for its role
in cell cycle arrest and apoptosis signalling (Ko & Prives, 1996). P53 acts as a transcription factor
for genes involved in apoptosis and cell cycle arrest, thereby nuclear translocation of the protein
is essential for its function (Ko & Prives, 1996). Under normal circumstances, p53 has been
observed in the cytoplasm of bovine blastocysts (Matwee et al., 2000), and its translocation to the
nucleus may be regulated by HSPs. In the preimplantation embryo, p53 is expressed at low levels
(Li et al., 2005). Stress experienced by the preimplantation embryo may result in an upregulation
of p53 as well as nuclear localization of its protein (Li et al., 2005).
The role of p53 during embryonic development is controversial. P53 may not play a role
25
in embryonic developmental arrest (Favetta et al., 2004), and apoptosis under non-stressed
conditions has been suggested to occur independently of p53 (Matwee et al., 2000). However, the
role of p53 appears to be critical in the embryonic response to environmental stress (Lichnovsky
et al., 1998; Hu et al., 2011), though it was not upregulated in response to oxidative stress in bovine
embryos (Favetta et al., 2007). In times of stress, p53 induces apoptosis in embryos thus ridding
the embryo of cells containing DNA damage (Stewart & Pietenpol, 2001), and p53 null embryos
exhibited low apoptosis and a high proportion of developmental abnormalities (Nicol et al., 1995;
Norimura et al., 1996). Furthermore, it appears that the levels of p53 in the early embryo may be
essential for proper development (reviewed by Choi & Donehower, 1999). Over- or underexpression of p53 can lead to increased incidence of malformations or embryo death (reviewed by
Choi & Donehower, 1999). Levels of p53 mRNA in the blastocyst may therefore indicate whether
the embryo is experiencing undue stress and give insight into its developmental fate.
Another well documented stress commonly encountered in embryos produced in vitro is
oxidative stress. Oxidative stress results in lower developmental competence and blastocyst quality
in IVP embryos (Batt et al., 1991; Farrell & Foote, 1995; Thompson et al., 1990; Rho et al., 2007).
Concurrent with this decrease in embryo development rate and quality, is an altered expression of
GLUT1, VEGF, Bax, and Bcl-2 among others as a result of varying levels of oxygen tension in
vitro (Rho et al., 2007). For instance, GLUT1 expression decreased with increasing oxygen
tension, with in vivo embryos exhibiting the highest expression of GLUT1 (Batt et al., 1991).
Furthermore, downregulation of GLUT1 can lead to decreased blastocyst cell number
(Balasubramanian et al., 2007). However, higher levels of GLUT1 expression as a result of
oxidative stress were observed in a similar study (Rho et al., 2007). It was suggested that this
difference may be due to altered embryonic metabolism as a result of oxidative stress experienced
26
by the IVP embryos (Rho et al., 2007).
GLUT1 is expressed at all embryonic developmental stages (Lequarre et al., 1997;
Bertolini et al., 2002) and becomes increasingly important following compaction, when the
embryo begins to use glucose as its primary energy source (reviewed by Pantaleon et al., 2001;
Harvey et al., 2004). The embryonic cellular uptake of glucose is mediated by GLUT1, and
expression has been observed to increase as a result of cellular stress caused by glucose deprivation
(Baldwin, 1993). Similarly, culture environments can affect the levels of GLUT1 gene expression
in bovine embryos (Wrenzycki et al., 1999, 2001; Lazzari et al., 2002) indicating that embryos
may respond to environmental conditions by altering GLUT1 expression levels (Harvey et al.,
2004).
It is therefore apparent that genes that are involved in physiological processes such as
apoptosis and metabolism during early embryo development can be affected by the conditions to
which the embryo is exposed. However, the effects that conditions during oocyte maturation have
on blastocyst gene expression is controversial. Some reports have suggested that blastocyst gene
expression may be altered due to maturation conditions (Silva et al., 2013), whereas others have
not observed such effects (Knijn et al., 2000). Heat and oxidative stress are common stress
pathways studied in the early embryo, and knowledge of these pathways is important in
understanding how exogenous stresses may impact embryo development.
Bisphenol A
Bisphenol A (BPA; 4,4’isopropylidenediphenol) has garnered much attention over the last
few decades due to its endocrine-disrupting properties, particularly for its effects as an estrogen
agonist (Nishikawa et al., 2010). BPA is a synthetic chemical that is polymerized in order to
27
produce polycarbonate plastics, epoxy resins, and flame retardants (Pastva et al., 2001; Lahnsteiner
et al., 2005; Nishikawa et al., 2010). BPA is one of the highest volume synthetic chemicals
produced worldwide (Welshons et al., 2006; Pearce et al., 2009; Vandenberg et al., 2009), and the
yearly production of BPA continues to increase and is expected to exceed 5.4 million tons in 2015
(Merchant Research & Consulting, 2014).
A wide range of plastic products contain BPA, including toys, water pipes, electronics,
baby bottles, medical equipment, tubing, and dental sealants (Shelby, 2008; Aghajanova &
Giudice, 2011). Incomplete polymerization results in the leaching of BPA from epoxy resins that
are used to line the inner surface of metallic food cans (Vandenberg et al., 2009). BPA has also
been found to leach from polycarbonate plastics such as baby bottles and reusable water bottles,
resulting in ingestion of BPA (Vandenberg et al., 2009). Ingestion is the primary route of BPA
contamination, although there is also evidence for dermal and inhalation exposure (Mørck et al.,
2010). BPA levels in various populations around the globe have been tested and BPA has been
found to be present in 92.6% (Wetherhill et al., 2007) of Americans, and about 90% of Canadians
(Bushnik et al., 2010) and has been detected in human urine, blood, serum, follicular fluid, and
tissue samples (Ikezuki et al., 2002; Schönfelder et al., 2002; Calafat et al., 2005; Vandenberg et
al., 2007). Thus it is imperative that we understand how this chemical acts and what risks it poses
to us and future generations.
Metabolism and Concentration
There are many controversies regarding BPA exposure and the study of BPA. One of these
involves its metabolism. A large amount of BPA is metabolised by the liver, and to a lesser extent,
the intestine. BPA is primarily glucuronidated to BPA-G with the help of cytochrome p450. The
28
next most common conjugation of BPA is sulfonation to BPA-S. Conjugated forms of BPA are
considered to be biologically inactive, having lost their estrogenic potential (Matthews et al.,
2001), however there is evidence that the toxicity of certain BPA metabolites may be even more
potent than unconjugated BPA (Yoshihara et al., 2004; Ishibashi et al., 2005; Baker &
Chandsawangbhuwana, 2012).
BPA is not metabolised to the same extent in all individuals, species, and tissues. BPA
metabolism is thought to be significantly influenced by an individual's genotype. Furthermore,
there are sex and age differences in the ability to metabolise BPA, with that of infant and young
children being relatively inefficient (Beydoun et al., 2014; Takeuchi et al., 2004; Doerge et al.,
2011; Yang et al., 2013). The varied responses that different species exhibit to BPA also make it
difficult to study. For instance, the rate of metabolism in humans differs from that in mice and rats.
Some studies have suggested that BPA is metabolized more quickly in rats (Elsby et al., 2001)
whereas Pritchett et al. (2002) predicted that when metabolic levels, which were slowest in human
hepatocytes, were extrapolated to the whole liver, humans had a greater capacity for BPA
metabolism than rats and mice.
Differences have also been found to occur at the tissue level. Much of BPA is conjugated
in the liver, and there is evidence of kidney and intestinal glucuronidation, however other tissues
are inefficient or unable to metabolise BPA (Trdan Lušin et al., 2012). The lungs, for instance, are
unable to metabolise BPA, making inhalation exposure of particular concern. There is also
evidence that some tissues are able to deconjugate BPA-G and BPA-S back into free BPA
(reviewed by Vandenberg et al., 2009). For instance, intestinal tissue can deconjugate BPA in the
presence of glucuronidases and sulfatases, releasing the bioactive form of BPA back into the tissue
(Zalko et al., 2003). Perhaps equally important as the metabolism is the presence of bound and free
29
BPA in human serum as well as in a wide variety of tissues (Calafat et al., 2005, 2008).
The concentration of BPA used in research is an additional factor contributing to the
complexity of its study. BPA has been found in a wide variety of serum and tissue samples however
controversy remains as to how much is actually in the tissue, and whether or not it is enough to
pose a risk to humans. The concentration of BPA detected in humans varies base on a person's age,
sex, race/ethnicity, household income or geographical location (Calafat et al., 2005, 2008). Levels
of free BPA in human serum has been found at levels ranging from 0.2 to 20 ng/mL with an
averages in the 1-3 ng/mL range in adults (Ikezuki et al., 2002; Vandenberg et al., 2007; Calafat
et al., 2008). Most relevant to the current study is the amount of free BPA found in follicular fluid.
Less data exists regarding follicular fluid content of BPA, but it has been found to average around
2.4 ng/mL in women undergoing IVF treatment (Ikezuki et al., 2002).
Since BPA can still be found in tissues and follicular fluid despite its relatively short halflife, it is thought that humans are exposed to significant amounts of BPA by frequent, low-dose
exposure through various sources (reviewed by Lenie et al., 2008). The concentrations measured
in human samples are much lower than the lowest-observed-adverse-effect-level (LOAEL) which
is considered to be 50 μg/mL as well as the in vitro culture equivalent which is suggested to be 50
ng/mL (Welshons et al., 2006; Wetherill et al., 2007). Of notable concern is that environmentally
relevant concentrations (low nanomolar range) have been found in many instances to result in
detrimental reproductive effects.
In addition to individual and tissue variability, exposure to BPA is not static, and these
concentrations will fluctuate in individuals depending on their eating habits and environmental
exposure levels. In a study by Carwile et al. (2011), it was found that ingestion of canned soup, in
comparison to fresh soup, resulted in an increase in urinary BPA levels in all volunteers tested.
30
Bpa has also been found to bind to human serum proteins (Csanády et al., 2002), including human
serum albumin (Yang et al., 2015). Therefore low concentrations of exposure may result in only a
small fraction of unbound BPA in plasma (Csanády et al., 2002). Thus, although exposure is
ubiquitous and constant, the actual concentrations found in humans are likely to vary. There are
many factors to consider when studying BPA, especially when using an animal model, and these
factors must be taken into account when interpreting results obtained in any study hoping to
extrapolate results to human exposure risk.
Known effects of BPA
BPA is an EDC best known for its ability to mimic estrogen, though it has been cited to
have many effects in various physiological systems. BPA toxicity is linked to the interference of
hormone regulation and to the disruption of the immune, reproductive, and neurological systems
(Rubin et al., 2001; Moriyama et al., 2002; MacLusky et al., 2005; Palanza et al., 2008; AvissarWhiting et al., 2010; Nishikawa et al., 2010). Of greatest relevance to the current work are the
effects BPA incurs on reproduction, particularly that of the oocyte and early embryo. The
reproductive effects of BPA published to date are vast. Altered hormone secretion (Vandenberg et
al., 2009), decreased implantation success (Berger et al., 2010; Ehrlich et al., 2012a), and altered
ovarian morphology (Suzuki et al., 2002) are a few of the many observed reproductive effects. Its
effects are also evident during early pregnancy with reports including altered early embryonic
development (Takai et al., 2001) and recurrent miscarriage (Kwintkiewicz et al., 2010).
31
BPA and the oocyte
BPA has been documented to affect both the prenatal and adult follicle and oocyte. Both
in vitro and in vivo studies have demonstrated BPA’s effects on the developing ovary. The onset
of meiosis, as well as GVBD and follicle formation have been demonstrated to be altered by
exposure to BPA (reviewed by Peretz et al., 2014). Some studies have found that the onset of
meiosis is affected by BPA exposure, resulting in meiotic nondisjunction in the fetal ovary without
evidence of aneuploidy (reviewed by Richter et al., 2007); however, aneuploidy has been observed
in other studies as a result of BPA exposure during early development. For instance, gestational
exposure during ovary development resulted in meiosis disruption, with fetal oocytes displaying
gross aberrations, leading to increased aneuploidy in oocytes and embryos of the mature female
(Susiarjo et al., 2007). Thus BPA may disrupt oogenesis in the developing ovary as a result of
maternal exposure. Similar effects have been observed as a result of BPA exposure during oocyte
maturation in the adult female, and these effects have been observed in humans and mice via in
vitro and in vivo studies.
Negative associations have been found between BPA serum levels and the number of
oocytes retrieved, peak serum E2 levels, the number of mature oocytes retrieved, oocyte
fertilizability, and oocyte developmental potential in women undergoing IVF (Mok-Lin et al.,
2010; Ehrlich et al., 2012b; Bloom et al., 2011a; Fujimoto et al., 2011). BPA exposure during
oocyte maturation in mammalian studies has resulted in meiotic abnormalities such as delayed cell
cycle progression (Can et al., 2005), spindle aberrations (Can et al., 2005; Eichenlaub-Ritter et al.,
2008), misalignment of chromosomes (Hunt et al., 2003; Eichenlaub-Ritter et al., 2008),
centrosomal alterations (Can et al., 2005) and increased aneuploidy (Hunt et al., 2003; Susiarjo et
al., 2007), all of which are indicators of poor oocyte quality. These effects have been shown to
32
vary with different exposure levels. Higher doses of BPA are linked to decreases in meiotic
progression, and lower doses resulting in increased abnormalities of MII oocytes (Lenie et al.,
2008; Machtinger et al., 2013). These data provide strong evidence that BPA at certain relevant
exposure levels may detrimentally affect oocyte maturation.
BPA and the embryo
Despite the many reported effects of BPA on female fertility and reproduction, the possible
effects of BPA on preimplantation development in mammals is not as well documented. There is
evidence that gestational or perinatal exposure can result in short- and long-term effects with the
possibility of a grand-maternal effect. Developmental exposure to BPA has resulted in poor
pregnancy outcomes such as pre-term birth (Cantonwine et al., 2010), predisposition to the
development of metabolic syndrome (Wei et al., 2011) as well as alterations in genes and
behaviour of offspring that may be passed on to the subsequent generation (Wolstenholme et al.,
2012). However, fewer studies have analyzed the effects of early exposure to BPA on blastocyst
development and quality in mammals. Existing studies have found detrimental outcomes as a result
of preimplantation BPA exposure with evidence suggesting that embryo development may be
altered as a result of BPA exposure.
In humans, urinary BPA concentration in women undergoing IVF was negatively
associated with blastocyst formation, but not embryo quality (Ehrlich et al., 2012b). Additionally,
male but not female urine BPA concentrations were found to decrease embryo quality in a
prospective cohort study of couples undergoing IVF (Bloom et al., 2011b). In vivo animal models
have demonstrated varying maternal BPA exposure doses can alter embryo development (Tsutsui
et al., 1998; Xiao et al., 2011). Despite delayed development, there was no alteration in sex ratio
33
of pups at weaning (Xiao et al., 2011). Conversely, Yan et al. (2013) found no differences in
development rate or total blastocyst number, but decreased hatching rates and increased proportion
of apoptotic cells were observed.
In vitro studies examining BPA exposure during preimplantation development have
exhibited similar results, and the same variability, as in vivo studies. Takai et al. (2000, 2001)
observed opposite effects of low (1-3 nM) and high (100 μM) doses of BPA on murine embryos.
Two-cell embryos exposed to 100 μM resulted in fewer embryos reaching the blastocyst stage,
whereas treatment with 1nM BPA resulted in a great number of embryos reaching blast. Embryo
quality and sex ratio did not differ between treatment groups and the control (Takai et al., 2000).
Additionally, both the high and low treatments led to pups that were significantly heavier than
controls at weaning (Takai et al., 2001). Thus BPA exposure during the preimplantation period
has the potential to affect blastocyst development (Xiao et al., 2011), as well as induce postnatal
effects. More information is needed regarding early exposure to BPA and developmental effects
on the early embryo as well as blastocyst quality in mammals.
BPA and male reproductive effects
Human studies regarding the male effects of BPA on sperm quality are limited but
generally agree that higher urinary BPA levels in infertile, but not fertile, men are associated with
decreased sperm count and motility (Mendiola et al., 2010; Li et al., 2011). Male urinary BPA
concentrations have also been associated with low embryo quality in IVF produced embryos
(Bloom et al., 2011b). Furthermore, prenatal or early postnatal exposure had resulted in adverse
effects in adult spermatogenesis and sperm quality of rodents (reviewed by Richter et al., 2007;
reviewed by Peretz et al., 2014). Exposure to low-dose BPA during gestation has been reported to
34
decrease sperm count in rats (Salian et al., 2009) as well as the number of elongated spermatids in
seminiferous tubules of pubertal mice (Okada & Kai, 2008). In addition to developmental
exposures, adult exposure has resulted in decreased sperm counts and increased apoptosis in rats
(Jin et al., 2013; Tiwari & Vanage, 2013). Additionally, sperm motility in rats and mice has been
impaired as a result of low-dose BPA exposure through various exposure routes as well as with
various developmental, and adult, exposures (Salian et al. 2009; Minamiyama et al. 2010;
Dobrzynska and Radzikowska 2013). Thus although the current thesis does not examine the effects
of male exposure to BPA, implications of male exposure on embryo development are important to
consider when interpolating in vitro results into real world significance.
BPA and gene expression
Gene expression analyses are commonly utilized to determine differences between
conditions and treatments of embryos developed in vitro, and is an excellent tool to examine the
mechanistic action of BPA. BPA has been reported to alter the expression of a variety of genes in
different cell and tissue types. Hormone receptors (Rubin, 2011), cell cycle regulators (Peretz et
al., 2012), apoptotic genes (Peretz et al., 2012), genes related to the stress response (Tabuchi et al.,
2002), and those related to the onset of meiosis, chromatin modification, remodeling, and
chromosome condensation (Lawson et al., 2011), among others have been altered as a result of
BPA exposure. Furthermore, BPA has been shown to exhibit nongenomic effects (reviewed by
Vandenberg et al., 2009), decrease methylation (Dolinoy et al., 2007), and alter imprinted gene
expression (Susiarjo et al., 2013). Analysis of how BPA affects gene expression, as well as
epigenomic responses, is important in the determination of how BPA exerts its action as a result
of early exposure.
35
Nuclear receptors
BPA has been documented to bind both the ERα and ERβ, though its influence on gene
transcription and its reproductive effects in the literature varies (Matthews et al., 2001; Kang et
al., 2006; Takao et al., 2003; Levy et al., 2004; Lahnsteiner et al., 2005). As mentioned, the primary
ligand of the ERs is E2. BPA competes with E2 to bind with the ERs (Levy et al., 2004), and it is
thought that some of BPA’s effects are elicited by its binding to these receptors (Welshons et al.,
2006; Chapin et al., 2008; Aghajanova & Giudice, 2011). BPA has been found to alter ERα and
ERβ under different experimental conditions, however BPA has higher affinity for ERβ than for
ERα (Matthews et al., 2001; Vandenberg et al., 2009). Additionally, BPA’s affinity for the ERs in
relation to E2 is weak, and reports of BPA’s estrogenicity in the literature vary (reviewed by Berger
et al., 2010).
Nonetheless, BPA has been found to alter gene and protein expression of ERs in a number
of reproductive (and non-reproductive) tissues. BPA exposure has resulted in altered ER
expression in vivo and in vitro. Observed effects include upregulation of ERα in uterine epithelium
(Markey et al., 2005), increased ERα and ERβ in brain tissue (Ramos et al., 2003), and increased
ER protein expression in uterine cells following a non-monotonic dose response (Berger et al.,
2010). In vitro BPA exposure has resulted in downregulation of ERα in sertoli cells (Tabuchi et
al., 2002) and endometrial stromal cells (Aghajanova & Giudice, 2011). However, as the ER
antagonist, ICI, has no effect on endometrial stromal cell gene expression, the effects of BPA may
not be mediated by the ERs (Aghajanova & Giudice, 2011).
Susiarjo et al. (2007) observed that meiotic defects as a result of maternal BPA exposure
resembled that of ERβ knockout mice. Additionally, ERβ knockout mice did not exhibit any
further BPA effects, suggesting both that BPA acts via the ERβ to exert its effects on fetal oocytes,
36
and fetal oocytes are sensitive to estrogenic actions. Preimplanation development can also be
influenced by BPA exposure via the ER (Takai et al., 2000). Culture of 2-cell mouse embryos in
vitro exposed to BPA resulted in dose-dependent effects on embryo developmental rates and
success (Takai et al., 2000). Exposure to 1 nM and 3 nM BPA resulted in increased development
rate whereas 100 μM BPA decreased the proportion of embryos developing to blastocyst. These
effects were inhibited by co-exposure to the anti-estrogen tamoxifen, suggesting BPA may be
acting via the ERs.
Although much attention has been paid to the estrogenic actions of BPA, interactions are
also known to exist between BPA and the TRs, antagonizing TH actions both in vitro and in vivo
(Moriyama et al., 2002; Iwamuro et al., 2003; Zoeller et al., 2005; Vandenberg et al., 2009; Meeker
& Ferguson, 2011). Activation of the TR by T3 is inhibited by BPA, resulting in antagonistic
effects on TR-response genes (Moriyama et al., 2002). BPA has been found to inhibit T3 action by
reducing T3 binding to the TR, suppressing transcriptional activities that are mediated by the TRs,
and recruiting nuclear receptor corepressor (N-CoR) to the TR promoter, resulting in
transcriptional inhibition (Moriyama et al., 2002). BPA is able to suppress TR-mediated
transcription by inhibiting positively regulated genes and by activating negatively regulated genes
(Moriyama et al., 2002; Heimeier et al., 2009). Although T3 is studied more widely due to its
prevalence compared to T4, BPA has also been found to exhibit an inverse relationship with T4
(Meeker & Ferguson, 2011).
Due to the importance of THs to preimplantation development, the effects of chemicals
such as BPA which have been reported to disrupt thyroid signalling, should be assessed in the
early embryo. In fish models low level BPA exposure during early development accelerated
embryonic development and time to hatch (Ramakrishnan & Wayne, 2008) and decreased growth
37
(Ramakrishnan & Wayne, 2008; Aluru et al., 2010). Admiodarone, a TR-antagonist blocked the
alterations in body growth and time to hatch in medaka fish (Oryzias latipes) (Ramakrishnan &
Wayne, 2008), suggesting that BPA may act through a thyroid pathway to disrupt early
development. However, it has been suggested that thyroid effects of BPA may require higher doses
of BPA than is required to produce estrogenic or antiestrogenic actions (Welshons et al., 2003).
Stress and metabolism
In addition to disruption of hormone pathways, BPA has been reported to alter gene
expression related to stress pathways and metabolism. For instance, genes which have been
observed to be overexpressed as a result of stress (such as oxidative or endoplasmic reticulum
stress) in mammalian cells were also upregulated following BPA exposure (Tabuchi et al., 2002).
These results suggest that BPA can induce cellular stress (Tabuchi et al., 2002). Additionally, BPA
has been found to decrease antioxidant enzymes in rat (Chitra et al., 2003) and mouse tissues
(Kabuto et al., 2003).
Apoptosis, which can be used to evaluate cellular stress, has been found to be induced by
BPA in various in vitro and in vivo models. It has been postulated that BPA may induce apoptosis
in cells by upregulating the proapoptotic genes Bax and p53 in rat embryonic midbrain cells in
vitro (Liu et al., 2013), as well as in murine antral follicles following maternal exposure to BPA
(Peretz et al., 2013). As discussed earlier, p53 can regulate apoptosis induction by activating
transcription of various proapoptotic genes (Amaral et al., 2010), thus increased p53 expression as
a result of BPA exposure may indicate a mechanism through which BPA exerts cellular stress.
Furthermore, BPA has been shown to upregulate hsp70 indicating increased stress experienced in
species of worm (Schirling et al., 2006), dinoflagellate (Guo et al., 2012), insect larvae (Planelló
38
et al., 2008), and crab (Park & Kwak, 2013). Further information on whether BPA influences
hsp70 transcription in mammalian embryos is required.
The metabolic effects of BPA during development have been studied in terms of rate of
development, and the relationship between prenatal BPA exposure and metabolic disorders later
in life. As mentioned, exposure of 2-cell embryos resulted in accelerated development at low
exposure levels, and delayed development at higher exposure levels, and these results were
reversed by tamoxifen supplementation (Takai et al., 2000). Furthermore, embryonic exposure to
various levels of BPA resulted in significantly increased weight of pups at weaning (Howdeshell
et al., 1999; Takai et al., 2000, 2001; Rubin et al., 2001; Markey et al., 2003; Akingbemi et al.,
2004). That developmental BPA exposure can result in altered metabolism has been observed in
various analyses. Perinatal exposure to BPA at the LOAEL dose led to increased body weight,
elevated serum insulin, impaired glucose tolerance, and decreased insulin sensitivity, all of which
were exacerbated by a high fat diet leading to metabolic syndrome (Alonso-Magdalena et al., 2010;
Wei et al., 2011).
At the molecular level, BPA exposure of adipocytes in vitro resulted in an upregulation of
mRNA of glucose transporters GLUT1 in human (Valentino et al., 2013) and GLUT4 in mouse
(Sakurai et al., 2004) as well as glucose uptake and utilization in both cases. Information regarding
BPA’s effects on gene expression, as well as the stress and metabolic responses in the early embryo
is limited. Considering the existing links between BPA and altered gene expression relating to
these pathways, the important roles of these genes in stress and metabolism, and the critical nature
of the embryonic response to stress, possible effects of BPA exposure on preimplantation stress
and metabolism is warranted.
39
RATIONALE, HYPOTHESIS, AND OBJECTIVES
Rationale
It is evident that BPA's influence is complicated and multifaceted, and it is clear why
reaching a consensus in regards to its mode of action has proven to be so difficult. Early
exposure to BPA can lead to a variety of detrimental alterations in development, throughout life,
and, as recent evidence suggests, onto future generations. Reproductive ability is compromised
by early BPA exposure by decreased gonad viability and fertility, and the ability of the embryo
to implant in the uterus. Those embryos that do successfully implant are exposed directly to BPA
through placental transfer, which can have serious implications on development by altering
serum levels, mRNA expression, and synthesis of various hormones and cytokines. This may
explain BPA's relationship to a plethora of diseases related to disruption of the reproductive,
immunological, and/or neurological systems.
Although the biological effects of BPA have been widely studied in recent years, studies
concerning embryo development in mammals are sparse and consist largely of embryonic
exposures. We know that BPA is present in the follicular fluid, and thus must also examine the
potential effects of BPA exposure at the oocyte stage on oocyte competence and quality of the
resulting blastocyst. A full picture of exposure during both the oocyte and embryo stages is
required, but first exposure during each of these stages separately must be assessed to get a
greater idea of how exposure at various steps affects early development. Studies that have
examined oocyte exposure have looked at effects on the oocyte itself but not later embryonic
effects. The purpose of the current research is to determine the effects of BPA exposure during
oocyte maturation on both the oocyte and embryos arising from these oocytes.
40
Hypothesis
The hypothesis of this study is as follows: exposure of bovine oocytes during in vitro
oocyte maturation to BPA will disrupt oocyte maturation, decreasing oocyte quality and
developmental potential, and compromising preimplantation embryonic development.
Objectives
To test this hypothesis, the following objectives were addressed:
Objective 1: To determine the effects of BPA exposure on oocyte maturation and
resulting quality.
Objective 2: To determine the effects of BPA exposure during oocyte maturation on early
embryo development.
Objective 3: To identify the effects of BPA exposure during oocyte maturation on mRNA
levels of key target genes in MII oocytes and blastocysts.
41
CHAPTER ONE*
Bisphenol A exposure during oocyte maturation in vitro results in spindle abnormalities
and chromosome misalignment in Bos taurus
*A portion of the material in this chapter has been published in Cytogenetics and Genome
Research (Ferris et al., 2015), and is reproduced with the permission of S. Karger AG, Basel,
Switzerland
42
INTRODUCTION
Bisphenol A (BPA) is an endocrine-disrupting chemical (EDC) used in the manufacturing
of many products consisting of polycarbonate plastics and epoxy resins. It is perhaps best studied
for its estrogenic actions and its ability to interact with estrogen receptor beta (ERβ) (reviewed by
Mtango et al., 2008). Many reproductive effects of BPA have been cited, yet controversy remains
concerning its reproductive toxicity. BPA has been detected in blood, serum, urine, and tissue
samples, and has been found in follicular fluid at an average concentration of 2.4 ng/mL in women
(Ikezuki et al., 2002). Humans are thought to be exposed to significant amounts of BPA by
frequent, low-dose exposure through various sources (reviewed by Lenie et al., 2008). Exposure
to BPA primarily occurs as a result of incomplete polymerization leading to the leaching of BPA
from epoxy resins that are used to line the inner surface of metallic food cans as well as from
polycarbonate plastics such as baby bottles and reusable water bottles (Vandenberg et al., 2009).
Ingestion is the primary route of BPA contamination, although there is also evidence for dermal
and inhalation exposure (reviewed in Mørck et al., 2010).
Negative associations have been found between BPA levels in urine or serum and number
of oocytes retrieved, peak E2 levels, normal oocyte fertilization and oocyte developmental
potential in women undergoing in vitro fertilization (IVF) (Mok-Lin et al., 2010; Fujimoto et al.,
2011). BPA exposure during oocyte maturation in mammalian studies has resulted in meiotic
abnormalities such as delayed cell cycle progression (Can et al., 2005), spindle aberrations (Can
et al., 2005; Eichenlaub-Ritter et al., 2008), misalignment of chromosomes (Hunt et al., 2003;
Eichenlaub-Ritter et al., 2008), centrosomal alterations (Can et al., 2005) and increased aneuploidy
(Hunt et al., 2003; Susiarjo et al., 2007), all of which are indicators of poor oocyte quality. These
43
data provide strong evidence that BPA at certain relevant exposure levels may detrimentally affect
oocyte maturation.
Oocyte maturation is a critical period of oocyte development that can determine the
developmental potential of the oocyte. Meiosis progression, spindle morphology, and chromosome
alignment at the metaphase plate are used as indicators of oocyte quality due to their importance
regarding further developmental success. The metaphase II (MII) spindle is often studied as an
indicator of oocyte quality due to its importance in the completion of meiosis, polar body extrusion
and segregation of the chromosomes. An association has been found in mammalian models
between spindle abnormalities and aneuploidy incidence which may result in pregnancy loss or
genetic diseases (Rama Raju et al., 2007; Ye et al., 2007; Tomari et al., 2011).
Poor quality oocytes are unable to properly support embryo development, and
developmental outcomes are significantly affected by the quality of the oocyte from which they
developed (reviewed by Mtango et al., 2008). External factors that have the potential to interfere
with normal oocyte maturation must therefore be thoroughly assessed in order to understand the
potential risk. The current study was designed to evaluate the effects of BPA during bovine in vitro
oocyte maturation on meiosis progression, as well as spindle formation and chromosome
alignment in MII oocytes. Based on findings in the literature, we hypothesized that exposing
bovine oocytes to BPA during oocyte maturation would result in decreased meiosis progression
and increased spindle abnormalities, thereby decreasing oocyte quality.
44
MATERIALS AND METHODS
Experimental design
Oocytes were matured in vitro (described below) in one of five treatment groups: (1) notreatment control (IVM), (2) vehicle control (0.1% ethanol), (3) E2 (2 μg/mL), (4) 15 ng/mL BPA
(65 nM), and (5) 30 ng/mL BPA (130 nM). The concentrations of BPA used in this study are
within the range that have previously been shown to induce MII abnormalities in mouse (Lenie et
al., 2008) and human (Machtinger et al., 2013) studies. Both concentrations fall below the
estimated lowest observed adverse effect level (LOAEL) exposure dose for in vitro cell studies
(Wetherill et al., 2007). These doses are higher than what has been measured in vivo since we
expect some BPA to bind to the plastic dish, be absorbed by the oil surrounding the IVM droplets,
and bind to bovine serum albumin present in the IVM media.
Chemicals
All chemicals were obtained from Sigma Life Sciences, Oakville, ON unless otherwise
stated.
Oocyte collection and in vitro oocyte maturation
Ovaries from domestic cattle (Bos taurus) were collected from a government inspected
abattoir (Cargill Meat Solutions, Guelph, ON, Canada). Cumulus oocyte complexes (COCs) were
aspirated from visible ovarian follicles for collection into HEPES-buffered Ham's F-10 plus 2%
steer serum (Cansera; Rexdale, ON, Canada). COCs containing several layers of cumulus cells
were collected and randomly assigned into one of five treatment groups. Groups of 10-15 COCs
were matured in 80 μL drops for in vitro maturation under silicone oil (Paisley Products, Toronto,
ON, Canada) for 24 hours at 38.5°C in 5% CO2 in air. The IVM drops consist of TCM199 medium
+ 2% steer serum supplemented with 1 μg/mL of E2, 0.5 μg/mL of bFSH and 1 μg/mL of bLH
45
(NIH, Washington, DC, USA). The treatment groups were composed of IVM media supplemented
with 0.1% ethanol, +1 μg/mL (to a total of 2 μg/mL) E2, 15 ng/mL BPA, or 30 ng/mL BPA.
Mature oocytes were stripped of their cumulus cells by gentle pipette-vortexing in
hyaluronidase (2 mg/mL in HEPES/Sperm TALP). Stripped oocytes were then washed 2 times in
PBS containing 0.1% polyvinyl alcohol (PBS-PVA) and either processed for ELISA (described
below) or fixed in 4% paraformaldehyde for 20 minutes at room temperature for
immunocytochemistry. Fixed oocytes were stored in PBS-PVA for no longer than 2 weeks at 4ºC
prior to staining.
Enzyme-linked Immunosorbent Assay (ELISA)
BPA concentrations were measured using Bisphenol A ELISA Kits (Creative Diagnostics,
DEIA12664). This kit can measure BPA levels in various fluids and cells using the procedures
described below. It is a competitive ELISA kit by which BPA can be measured in samples through
a limited number of binding sites from the anti-BPA antibody which coat the bottom of the wells
of the ELISA plate. BPA epitope in the samples compete with BPA-HRP conjugate which is added
to all wells with the exception of the “blank” wells. Therefore the amount of BPA conjugate bound
to each of the wells is inversely proportional to the BPA concentration in the sample, and is
determined by the amount of color obtained when TMB, which reacts with the unbound HRP in
the well, is added. Sulfuric acid is used as a stop solution converting the blue coloured product
obtained with the addition of TMB to a yellow coloured product which can then be read on a plate
reader at 450 nm. The specificity of the BPA ELISA as reported by the manufacturer with regards
to BPA and related chemicals is as follows: BPA: 100%; BPS: <0.01%; Reversatrol: <0.01%.
46
IVM media used for all groups were processed immediately (T0), and at 24 hours following
incubation with maturing oocytes (T24+). Mature oocytes were removed from media samples prior
to processing the T24+ group. Acetic acid (Fisher Scientific Canada, Ottawa, ON, Canada) was
added to 1 mL media to a pH of 4. The following was repeated 3 times to yield 3 mL ethyl acetate
product. Equal parts ethyl acetate (Caledon Laboratories Ltd., Georgetown, ON, Canada) was
added to the media sample which was then vortexed for 20 seconds and centrifuged at 2000 RPM
for 10 minutes at room temperature for phase separation. The top (ethyl acetate) phase was
removed and placed in a glass culture tube. The bottom phase was removed with a glass pipette
and placed in a new tube for the extraction process to be repeated. Following three extractions, the
ethyl acetate was evaporated under N2 gas. Dried samples were then sealed and stored at -20°C
until use.
Oocytes for BPA concentration analysis were collected as follows: COCs were aspirated
from ovaries, washed in IVM without addition of hormones, and matured in treatment groups
containing 0.1% ethanol, 15 ng/mL BPA, or 30 ng/mL BPA, as described, for 24 hours. Immature
or mature oocytes were placed in 2 mg/mL hyaluronidase solution and gently agitated to remove
cumulus cells. Denuded oocytes were pooled into 3 biological replicates of 50 oocytes each and
placed in 0.5 mL DI water in a 1.5 mL eppendorf tube. Oocytes were sonicated for 15 seconds at
output power 4 using a Microson Ultrasonic Cell Disruptor (Misonix Inc., Farmingdale, NY,
USA). Acetic acid was added to a pH of 4. The following was repeated 3 times to yield 1.5 mL
ethyl acetate product. A volume of 0.5 mL ethyl acetate was added to each sample which were
vortexed for 1 min followed by centrifugation at 10,000 RPM for 5 minutes at room temperature.
Following centrifugation, two phases were visible. The top layer was removed and placed in a
47
glass culture tube, and the bottom was used to repeat the extraction procedure. Following the three
extractions, ethyl acetate was evaporated under N2 gas and stored sealed at -20°C.
ELISAs were performed according to the manufacturer’s instructions (Creative
Diagnostics, DEIA12664). Briefly, samples were thawed, reconstituted with 10 μL ethanol, and
diluted with 500 μL sample dilution buffer. Following addition of samples, standards, and HRP
conjugate, the ELISA plate was incubated at room temperature for 2 hours. The plate was washed
3x with wash buffer and dried by blotting on paper towel. TMB (provided with kit) was added to
each well (200 μL) and the plate was incubated 30 minutes at room temperature in the dark.
Sulfuric acid was added to each well (50 μL) as a stop solution and the plate was read with BioTek EL800 Universal Microplate Reader (Bio-tek Instruments Inc., Winooski, VT, USA) at 450
nm.
Immunocytochemistry and Imaging
Oocytes were stained to visualize microtubules and chromatin. Briefly, fixed oocytes
(described above) were permeabilized in 0.5% Triton-X in TBST for 90 minutes followed by 3
10-minute washes. Oocytes were blocked in 1% goat serum (Millipore Canada, Etobicoke, ON,
Canada) for 1 hour and incubated overnight at 4°C in 1:500 monoclonal anti-α-tubulin produced
in mouse (Sigma-Aldrich, St. Louis, MO, USA). The following day, oocytes were washed four
times (2x 30min, 2x 15min) in TBST and incubated in the dark overnight at 4°C in 1:500 secondary
antibody (Alexa-Fluor 488 Goat Anti-Mouse, Life Technologies, Burlington, ON, Canada).
Following the incubation, oocytes were washed three times for 30 minutes each in TBST and
transferred to a PBS wash.
Oocytes were mounted onto slides and PBS was evaporated by air drying to allow adhesion
to the slide. Vectashield containing DAPI (1.5 μg/mL) was placed on top of oocytes as a
48
chromosome stain. A syringe was used to place drops of Vaseline on the slide to avoid flattening
of the oocytes when covered with a coverslip. Slides were sealed with nail polish and stored in the
dark at 4°C for no longer than 2 weeks. Oocytes were imaged at 60x under oil using an Olympus
FV1200 Confocal Microscope with laser wavelengths of 405 nm for DAPI and 488 nm for AlexaFluor 488 using Fluoview software.
Analysis of Oocytes
Oocytes in all treatment groups were analysed for meiotic stage. Representative images are
shown in Figure 3. To calculate maturation success, the proportion of oocytes to reach MII versus
the proportion of oocytes that did not reach maturation was compared between each of the
treatment groups (Fisher’s exact; two-tailed).
Spindle formation and chromosome alignment analyses
Oocytes exhibiting a polar body and MII spindle were analyzed for spindle formation and
chromosome alignment. Seven oocytes were excluded from the spindle formation analysis due to
poor spindle view. Spindles were categorized as displaying a normal or abnormal spindle
formation. Spindles were considered normal when there were two poles without flattening,
equidistant from the metaphase plate (see Fig. 5A for example). Spindles were considered
abnormal when one or both poles were flattened, there was no focussed polar region, and a
flattening of the spindle led to a reduction in size (see Fig. 5B for example).
Chromosome alignment at the metaphase plate was analyzed and oocytes were categorized
as having chromosomes that were aligned or dispersed at the metaphase plate. Aligned
chromosomes showed little to no deviation from the metaphase plate (see Fig. 5A for example),
whereas dispersed chromosomes were identified as having one or more chromosomes out of
alignment with substantial deviation from the metaphase plate (see Fig. 5C for example).
49
Statistical analyses
ELISA results were interpreted using Graphpad Prism 6 software. Differences in
concentrations of BPA in the media and oocyte analyses were compared between samples using
one-way ANOVA and Tukey’s multiple comparison test. A p value of <0.05 was used to establish
statistical significance.
Two-tailed Fisher’s exact test was used to calculate differences between the treatment
groups of the proportion of oocytes to reach maturation versus those that did not, as well as the
proportion of MII oocytes displaying a normal vs. abnormal spindle and those displaying aligned
vs. dispersed chromosomes. A p value of <0.05 was used to establish statistical significance. The
proportion of oocytes to arrest in the earlier stages of meiosis were analyzed with one-way
ANOVA, and a p value of <0.05 was used to establish statistical significance.
50
RESULTS
Oocyte uptake of BPA and IVM media concentrations
The concentration of BPA measured in MII oocytes are shown in Figure 1. Results are
presented as the measured BPA concentration (ng/mL) per oocyte. MII oocytes matured with IVM
supplemented with the ethanol vehicle had an average uptake of 1.41 ng/mL BPA, and those
matured with IVM supplemented with 15 ng/mL and 30 ng/mL BPA had an average uptake of
1.69 ng/mL and 2.48 ng/mL respectively. The concentration of BPA measured in oocytes exposed
to 30 ng/mL was significantly higher than the other groups (p<0.01).
BPA concentrations in treatment group media are presented in Figure 2. BPA concentration
in the 0.1% ethanol group did not differ from the IVM group. Media of the 30 ng/mL BPA group
had significantly higher levels of BPA, as expected, than the other groups analyzed (p<0.05). Both
BPA groups exhibited a significant decrease in BPA following the 24 hour incubation with COCs
(p<0.05).
Meiosis progression and MII spindle abnormalities
Representative images of meiotic stages in bovine oocytes are displayed in Figure 3. The
proportion of oocytes to reach the MII stage in each of the treatment groups is summarized in
Figure 4. The 0.1% ethanol group did not differ from the IVM group. A lower proportion of
oocytes exposed to 30 ng/mL BPA during oocyte maturation reached MII than oocytes in the IVM
group (57.4% (62/108) and 72.4% (71/98), respectively; (p<0.05)). The proportion of oocytes in
the 30 ng/mL BPA group that arrested in germinal vesicle breakdown, prometaphase I, or
metaphase I was almost twice that of the no-treatment control group (IVM), however this was not
statistically significant (26.9% (29/108) and 15.3% (15/98), respectively).
51
Figure 1. Mean BPA concentrations of oocytes following IVM in their respective treatment
groups. Concentration of BPA (mean ± SEM) in oocytes at 24 hours following oocyte maturation
in IVM media supplemented with 0.1% ethanol (n=150), 15 ng/mL BPA (n=150), or 30 ng/mL
BPA (n=150). Analysis of variance (ANOVA) and Tukey’s multiple comparison test, **p<0.01.
52
Figure 2. Mean BPA concentrations of IVM media before and after incubation with oocytes.
Concentration of BPA (mean ± SEM) in IVM media with no supplementation (IVM), or
supplemented with 0.1% ethanol, 15 ng/mL BPA, or 30 ng/mL BPA prior to (T0) or at 24 hours
following oocyte maturation (T24+). Oocytes were removed prior to processing of T24+ media.
White asterisk = differences between T0 media samples of different treatment. Black asterisks =
differences between T0 and T24+ media samples of the same treatment. (Analysis of variance
(ANOVA) and Tukey’s multiple comparison test, *p<0.05).
53
Figure 3. Representative images of meiotic stages during bovine oocyte maturation.
Representative images of bovine oocytes during maturation with α-tubulin in green and chromatin
in blue. Stages include (A) germinal vesicle; (B) germinal vesicle breakdown; (C) prometaphase
I; (D) metaphase I; (E) anaphase I; (F) telophase I; (G) prometaphase II; (H) metaphase II with
polar body (arrow) visible. Scale bar: 20 = μm.
54
Figure 4. Proportion of oocytes to reach MII following IVM in their respective treatment
groups. Proportion of oocytes (mean ± SEM) to reach the MII stage following 24 hour incubation
in IVM media without (IVM; n=98) or with 0.1% ethanol (n=97), 15 ng/mL BPA (n=110), or 30
ng/mL BPA (n=108) supplementation. Fisher’s exact, *p<0.05.
55
MII oocytes in all groups were analyzed for spindle morphology and chromosome
alignment. Representative images are displayed in Figure 5. Figure 5B and C indicate spindle
abnormalities observed in oocytes exposed to 30 ng/mL BPA during maturation. Abnormal spindle
morphology and chromosome misalignment were frequently observed in oocytes which were
matured with 30 ng/mL BPA supplementation, with abnormal spindle morphology slightly more
prevalent than chromosome misalignment (Figs. 6 and 7). The spindle morphological abnormality
most frequently observed was of a compressed spindle and loss of focussed polar regions. In some
cases only one half of the spindle exhibited this abnormality, however most cases of abnormal
morphology consisted of significant microtubule compression at both spindle ends.
Figure 6 shows the proportion of MII oocytes that displayed abnormal spindle morphology.
Exposure to 30 ng/mL BPA during oocyte maturation resulted in a significantly higher proportion
of oocytes displaying abnormal spindle morphology compared to all of the other treatment groups
(67.9% (19/28); p<0.05). There were no significant differences between the remaining treatment
groups (IVM = 28% (7/25); 0.1% ethanol = 19% (4/21), 2 μg/mL E2 = 35.5% (11/31), 15 ng/mL
BPA = 31.3% (10/32)).
The proportion of oocytes displaying chromosomal dispersal is shown in Figure 7. A higher
proportion of oocytes assigned to the 30 ng/mL BPA treatment group exhibited chromosome
dispersal at the metaphase plate compared to all of the other treatment groups (IVM = 19.2%
(5/26), 0.1% ethanol = 29% (4/21), 2 μg/mL E2 = 25.8% (8/31), 15 ng/mL BPA = 21.2 (7/33), 30
ng/mL BPA = 60% (18/31); p<0.01). There were no significant differences between the other
treatment groups, including the lower dose of BPA.
56
Figure 5. Representative classifications of MII oocytes for spindle morphology and
chromosome alignment. Representative images of bovine MII oocytes with α-tubulin in green
(left panels), chromatin in blue (middle panels) and merged images (right panels). Oocytes with
normal spindle morphology and chromosomes alignment (A), abnormal spindle morphology (B),
and chromosome dispersal (C) are displayed. Rows B and C represent spindle abnormalities
observed in oocytes treated with 30 ng/mL BPA. Visible polar body is marked with an arrow.
Scale bar = 20 μm.
57
Figure 6. Proportion of MII oocytes displaying normal and abnormal spindle morphology
following IVM in their respective treatment groups. Proportion of oocytes in each of the
treatment groups displaying abnormal spindle morphology following 24 hour incubation in IVM
media without (IVM; n=25) or with 0.1% ethanol (n=21), 15 ng/mL BPA (n=32), or 30 ng/mL
BPA (n=28) supplementation. Fisher’s exact, *p<0.05.
58
Figure 7. Proportion of MII oocytes displaying dispersed chromosomes at the metaphase
plate following IVM in their respective treatment groups. Proportion of oocytes in each of the
treatment groups displaying dispersed chromosomes at the metaphase plate following 24 hour
incubation in IVM media without (IVM; n=26) or with 0.1% ethanol (n=21), 15 ng/mL BPA
(n=33), or 30 ng/mL BPA (n=31) supplementation. Fisher’s exact, **p<0.01.
59
DISCUSSION
In this study we have observed that exposure to 30 ng/mL BPA during bovine oocyte
maturation in vitro results in an average oocyte uptake of 2.48 ng/mL BPA, as well as decreased
meiosis progression and increased incidence of spindle abnormalities, specifically abnormal
spindle morphology and chromosome alignment. Other studies have suggested that exposure to
BPA during oocyte maturation may induce meiotic abnormalities in mouse and human oocytes.
This was first shown by Hunt et al. (2003) when mice were inadvertently exposed to BPA via
damaged water bottles and cages, resulting in meiotic disturbances in mouse oocytes. This
correlation was then confirmed experimentally with oral administration of BPA (Hunt et al., 2003).
The link between BPA and meiotic disruption has since been observed in mice (Can et al., 2005;
Susiarjo et al., 2007; Lenie et al., 2008) and humans (Machtinger et al., 2013) with varying results
and experimental parameters. For instance, meiotic abnormalities were observed in mice and
humans as a result of BPA supplementation of culture media in which COCs were matured in vitro
(Can et al., 2005; Machtinger et al., 2013). In the case of the human oocytes, clinically discarded
oocytes with partial disruption to the cumulus cells were utilized (Machtinger et al., 2013). Culture
of individual follicles in vitro supplemented with varying concentrations of BPA have also induced
meiotic abnormalities (Lenie et al., 2008). Additionally, the onset of meiosis during fetal ovarian
development was disrupted, as was oocyte maturation in the adult, following in vivo exposure of
pregnant mice via implantation of BPA pellets (Susiargo et al., 2007).
The current study was designed to examine meiosis progression and MII spindle integrity
following exposure to BPA during in vitro maturation of bovine oocytes, with the intent of
examining the link between oocyte exposure to BPA and the quality of oocytes and resulting
embryos. The current study employs a bovine in vitro maturation model due to the high
60
physiological similarities existing between the bovine and human reproductive systems. Bovine in
vitro oocyte maturation is an excellent model to evaluate female reproductive toxicology due to
similarities in follicular dynamics and endocrine control between bovine and human (Beker van
Woudenberg et al., 2012). Presently, we examined the effects of BPA exposure during oocyte
maturation on bovine oocyte quality.
Oocyte uptake of BPA and IVM media concentrations
The amount of BPA taken up by the oocytes was much lower than the initial exposure
levels. Oocytes exposed to 15 ng/mL took up an average of 1.69 ng/mL BPA whereas those
exposed to 30 ng/mL BPA took up an average of 2.48 ng/mL BPA (Fig. 1). These levels are similar
to those that have been found in follicular fluid samples in women. Women undergoing IVF were
found to have 2.4 ± 0.8 ng/mL BPA in follicular fluid samples (Ikezuki et al., 2002). The average
1.69 ng/mL BPA taken up by oocytes exposed to 15 ng/mL is similar to the lower range found in
these women, whereas the 2.48 ng/mL BPA taken up by oocytes exposed to 30 ng/mL is similar
to the average exposure level in the follicular fluid of women analyzed (Ikezuki et al., 2002). The
exposures used in this study are of environmental significance, and the concentrations taken up by
the oocytes are similar to levels of BPA that have been measured in follicular fluid of women.
The oocyte analysis of BPA was conducted using pools of oocytes to ensure a detectable
level of BPA. The values obtained were then divided to obtain an average concentration per oocyte.
Therefore the values presented in Figure 1 are only an average of what is being taken up by the
oocytes and do not necessarily represent the amount of BPA taken up by each individual oocyte.
There is likely variability between individual oocytes regarding the amount of BPA taken up
within each biological replicate. This could partly explain how all oocytes exposed to the same
levels of BPA do not respond identically. The amount of BPA taken up by an oocyte is likely a
61
determining factor in the results observed. We therefore consider the amount of BPA taken up by
the oocyte to be of greater importance than the initial exposure dose. Thus BPA concentration in
follicular fluid as well as that of experimental supplementation in IVM media does not directly
correlate with the amount of BPA taken up by the oocyte. This may be a result of uptake of BPA
by the surrounding cumulus cells and/or granulosa cells within the follicle, as well as varying
uptake rates of oocytes. The specific mechanism of BPA uptake into the oocyte needs further
analysis to determine how BPA is taken up by the oocyte as well as what factors contribute to
differences in BPA uptake among oocytes.
Although BPA has been quantified in follicular fluid, to our knowledge the measurement
of BPA in oocytes themselves has only been reported once previously. Consistent with the current
results, Aluru et al., (2010) observed that only a small proportion (1.2% and 4.5%) of BPA was
detected in rainbow trout (Oncorhyncus mykiss) oocytes following 3 hours of exposure in
comparison to the t0 measurements of 32 and 417 ng/oocyte respectively. Interestingly, the lack
of cumulus cells in the fish model suggests the amount of BPA taken up by the oocyte is limited
by the rate at which this uptake occurs.
The number of cumulus cells surrounding the oocyte and the initial quality of the oocyte
prior to IVM may affect the amount of BPA that is taken up by the oocyte. Further studies could
analyze if these factors have an influence on the oocyte’s susceptibility to BPA exposure by
evaluating the amount of BPA taken up by oocytes which have been stripped of their cumulus cells
versus those that haven’t, as well as oocytes of higher versus lower initial quality which can be
estimated based on their cytoplasmic appearance (Wood & Wildte, 1997). Determining whether
compromised oocytes take up different levels of BPA than non-compromised oocytes would
provide important information on oocyte susceptibility to BPA. Oocytes of suboptimal quality that
62
may survive under normal conditions may be unable to withstand the effects of BPA exposure at
concentrations that good quality oocytes may be able to endure. Thus this information would be
valuable in the future to further evaluate the effects of BPA on oocyte maturation and assess the
risk of human exposure.
Meiosis progression and MII spindle abnormalities
Meiosis progression, spindle morphology and chromosome alignment of MII oocytes are
related to oocyte quality and developmental potential (Rama Raju et al., 2007; Ye et al., 2007;
Tomari et al., 2011). Abnormalities in spindle morphology and chromosome alignment at the
metaphase plate have been found to lead to embryonic aneuploidy as a result of maternal factors
such as advanced maternal age and obesity (Battaglia et al., 1996; Luzzo et al., 2012). External
factors such as inappropriate culture conditions have also been shown to induce meiotic
abnormalities, resulting in decreased oocyte quality (Wang et al., 2002). The external
environmental factor currently discussed, BPA, has been shown under certain conditions to lead
to spindle aberrations, misalignment of chromosomes, and aneuploidy as a result of oocyte
exposure (Hunt et al., 2003; Can et al., 2005; Eichenlaub-Ritter et al., 2008; Lenie et al., 2008;
Machtinger et al., 2013). However, these trends are not observed under all conditions, thus we
aimed to identify meiotic and spindle effects in the current experimental model.
We have shown that exposure to 30 ng/mL BPA during maturation which results in an
average oocyte concentration of 2.48 ng/mL BPA leads to reduced oocyte maturation success, and
increased incidence of spindle abnormalities. We observed an 8-15% reduction in the proportion
of oocytes reaching maturity in this group compared to the other treatment groups (Fig. 4). The 15
ng/mL BPA treatment resulting in an average oocyte concentration of 1.69 ng/mL BPA did not
show a similar decrease, suggesting that a higher level of oocyte BPA uptake is required to
63
decrease maturation success. These results are in agreement with studies conducted in mouse (Can
et al., 2005; Lenie et al., 2008) and human (Machtinger et al., 2013). In dose-dependent studies,
meiotic arrest was largely only seen in the higher doses examined (Can et al., 2005; Lenie et al.,
2008; Machtinger et al., 2013), however the oocyte concentration of BPA in these studies is not
known. Furthermore, meiosis progression in the 30 ng/mL BPA group was only significantly
decreased in comparison to the IVM group but not the 0.1% ethanol group, indicating there is a
slight but non-significant effect of the vehicle that is exacerbated by the higher dose of BPA.
Ethanol at higher concentrations than currently used has previously been shown to elicit a stress
response in porcine COCs (Lee et al., 2014a), resulting in increased expression of pro-apoptotic
genes in oocytes and cumulus cells. Thus it is possible that the ethanol vehicle currently used is
causing slight but not significant alterations in maturation of the oocyte and must be considered
when interpreting results. However effects observed in the spindle and chromosome analysis did
differ significantly between the 30 ng/mL BPA group and the 0.1% ethanol group.
In the current analysis, MII oocytes with a BPA concentration of 2.48 ng/mL exhibited a
32.4 – 48.9% increase in the occurrence of abnormal spindle morphology and a 31 – 40.8%
increase in the occurrence of chromosome dispersal compared to the other groups (Figs. 6 and 7,
respectively). These results are in agreement with studies conducted in mouse (Hunt et al., 2003;
Lenie et al., 2008) and human (Machtinger et al., 2013), which have found MII abnormalities as a
result of BPA exposure during oocyte maturation. However, the exposure levels of BPA, and likely
the resulting oocyte concentration of BPA, are critical to the results obtained, with exposure levels
and responses varying between studies. For instance, MII spindle abnormalities were observed
following exposure levels of 3 nM (0.685 ng/mL) to 3 μM (685 ng/mL), but not 30 μM (6.85
μg/mL), during mouse in vitro follicular development (Lenie et al., 2008). Additionally, a
64
significant increase in MII abnormalities were observed following BPA exposure to 20 ng/mL but
not 200 ng/mL during in vitro maturation of human oocytes (Machtinger et al., 2013).
Furthermore, Lenie et al. (2008) reported most MII abnormalities were due to misalignment of
chromosomes at the metaphase plate with relatively few abnormalities as a result of spindle
malformations, whereas in the current study, incidence of spindle malformation and chromosome
misalignment were similar, but there was a higher incidence of the oocytes containing an average
of 2.48 ng/mL BPA displaying both malformations (43.5%) rather than one or the other, compared
to the other groups (9.1- 23.1%).
Characteristics of the abnormalities also differ between studies. For instance, Hunt et al.
(2003) exposed mice in vivo to varying doses of BPA during folliculogenesis, which resulted in
spindle malformations, but the typical malformation was of an elongated spindle, in contrast to the
shortened spindle observed in our model. Chromosome dispersal observed in other studies was
also much more severe than found in the current analysis, with chromosomes in those studies
sometimes dispersed throughout the entire length of the spindle as has been observed in mouse
(Hunt et al., 2003; Lenie et al., 2008) and human oocytes (Machtinger et al., 2013). These
differences may be explained due to varying exposure lengths and concentrations, the time of
oocyte development at which exposure occurred, differences occurring between in vitro and in
vivo studies, the amount of BPA being taken up by the oocytes, as well as species specific
differences.
BPA has been shown to act in an estrogenic and estrogen-independent manner (reviewed
by Wetherill et al., 2007). The E2 group was included to compare results obtained following
additional E2 supplementation to IVM media with that of the two BPA treatments. E2 has
previously been shown to impair meiotic progression and spindle morphology in vitro (Beker et
65
al., 2002; Beker-van Woudenberg et al., 2004). In the current chapter, treatment with additional
E2 did not result in meiotic or spindle perturbations compared to the control groups. However, as
E2 is included in standard IVM protocols in our laboratory, the E2 group included additional E2
supplementation to a total of 2 μg/mL as has been done previously (Pocar et al., 2003), whereas in
the aforementioned studies, the control groups were void of E2 (Beker et al., 2002; Beker-van
Woudenberg et al., 2004). Thus it is possible that E2 is currently inducing meiotic abnormalities,
but the incidence of abnormalities do not differ between the additional E2 supplementation in the
E2 group, and the standard E2 treatment in the control groups. Additionally, Beker-van
Woudenberg et al. (2004) analyzed spindle perturbations throughout meiosis whereas we only
assessed the MII spindle for abnormalities. Additional analyses are required to determine the
mechanism(s) by which BPA is exerting its effects, and whether these may be through estrogenic
pathways. Interestingly, Beker-van Woudenberg et al. (2004) reported that the meiotic
abnormalities induced by E2 were not due to membrane receptor interaction.
The mechanism by which the meiotic spindle can be disrupted by BPA, however, is not
clear. It has been suggested that BPA may disrupt meiosis by direct interaction with the
microtubules and associated proteins during maturation (Pfeiffer et al., 1997; Can et al., 2005).
Selective interference with centrosome and microtubule organization by BPA may result in
meiotic disturbances observed (Can et al., 2005). BPA may induce errors in cell cycle progression
and microtubule assembly and function by targeting microtubule-associated motor proteins,
disrupting protein transport, or by interfering with protein interactions during oocyte maturation
(Takahashi et al., 2000; Can et al., 2005). Analyses of factors important to meiosis and spindle
assembly during oocyte meiosis is required to gain a greater understanding of how BPA may be
eliciting these effects.
66
As has been shown in studies linking decreased maturation success and developmental
outcome, meiosis progression and MII spindle morphology are indicators of oocyte quality. Thus
the main findings of this research suggest that oocyte quality is compromised following exposure
to 30 ng/mL BPA during maturation, resulting in an average oocyte concentration of 2.48 ng/mL
BPA. This is evidenced by decreased meiosis progression and increased spindle abnormalities and
chromosome dispersal at the metaphase plate. Considering the link between these indicators of
oocyte quality and future development, BPA exposure during oocyte maturation resulting in a
significant increase in oocyte concentration of BPA could have the potential to disrupt future
development of the oocyte. Further analysis is required to determine the factors influencing the
amount of BPA taken up by the oocyte, and potential embryonic effects of oocyte exposure to
BPA during maturation.
Exposure to 15 ng/mL BPA during in vitro maturation of bovine oocytes resulted in an
average uptake of 1.69 ng/mL BPA and showed no significant meiotic or MII effects against the
controls in the current analysis. However, the 30 ng/mL exposure level resulted in an average
uptake of 2.48 ng/mL BPA and did show significant effects in all parameters analyzed. Following
this exposure, fewer oocytes reached maturity, and a higher proportion of MII oocytes displayed
abnormal spindle morphology and chromosome dispersal at the metaphase plate. These factors are
important to oocyte quality and developmental potential, thus BPA’s ability to influence oocyte
maturation suggests that exposure during this time may affect embryonic viability.
67
CHAPTER TWO
Exposure to bisphenol A during in vitro oocyte maturation results in decreased embryo
development, skewed sex ratio, and increased apoptosis in blastocysts of Bos taurus
68
INTRODUCTION
Successful development of an early embryo will largely depend on its environment as well
as its innate ability to respond to environmental changes or stressors. Various conditions of stress
can result in consequences to the embryo including delayed or abnormal development, apoptosis
induction, altered gene expression, and miscarriage (Wilson et al., 1985; Niemann & Wrenzycki,
2000; Arck, 2004; Dennery et al., 2007; Ornoy, 2007). These stressors include hypoxia, changes
in temperature, pathogens, UV radiation, free radicals, environmental toxins, and poor maternal
diet or health (Betts & King, 2001; Paula-Lopes & Hansen, 2002b; reviewed by Hamdoun & Epel,
2007). The developmental fate of the embryo rests upon the embryo itself and its defense
capabilities of withstanding such stressors.
Apoptosis, or programmed cell death, eliminates abnormal or damaged cells as well as
cells with abnormal developmental potential (Hardy, 1999; Betts & King, 2001; Paula-Lopes &
Hansen, 2002a). In the blastocyst, cell death may be a way to regulate development, eliminating
abnormal or damaged cells (Betts & King; 2001, Hansen & Fear, 2011). By removing cells that
are damaged, abnormal, or have an inappropriate developmental potential, apoptosis acts as a
quality control mechanism and is an important factor in animal development (Jacobson et al., 1997;
Betts & King, 2001). The ability of the embryo to induce apoptosis in times of stress may help it
to survive suboptimal conditions, however the extent to which apoptosis occurs in the blastocyst
plays an important role in the fate of that embryo (Betts & King, 2001). Where mild induction of
apoptosis might help the embryo survive by eliminating damaged cells, severe stress may cause
extensive apoptosis that could compromise development and survival of the embryo (Byrne et al.,
1999; Betts & King, 2001; Paula-Lopes & Hansen, 2002b; Hansen & Fear, 2011).
69
Adverse environmental exposures during preimplantation development can increase the
proportion of cells undergoing apoptosis in the early embryo (Paula-Lopes & Hansen, 2002b).
Stresses encountered by preimplantation embryos include temperature change, heat stress,
oxidative stress, pathogens, and exposure to toxic or xenobiotic chemicals (Betts & King, 2001;
Hamdoun & Epel, 2007). These environmental stressors have been found to induce apoptosis in
early embryos under certain conditions and may compromise embryonic viability and survival
depending on the severity of the stress, and the time at which exposure occurs during development
(Paula-Lopes & Hansen, 2002a; Hamdoun & Epel, 2007). Therefore the fate of the embryo may
depend on the type of stress, degree of apoptotic response, and the time at which the embryo is
exposed to stress.
Another possible outcome of preimplantation stress is a skewed sex ratio. Maternal stress
has been found to impact the sex ratio of offspring in some cases. In rodents, females exposed to
social stress produced fewer sons in mice (Krackow, 1997), hamsters (Pratt & Lisk, 1989), and
rats (Lane & Hyde, 1973; Moriya et al., 1978), and the type of stress that may result in a decrease
in sex ratio (fewer males) includes but is not limited to stressors such as pollution (James, 1998)
and confinement stress (Krackow, 1997). Hyperglycemia-induced stress in murine embryos
resulted in both an increase in apoptosis as well as a skewed sex ratio in favour of females (Jiménez
et al., 2003), providing an explanation for the trend that more daughters are born to diabetic
mothers (Rjasanowski et al., 1998). Fewer males also tend to be conceived in humans under
suboptimal conditions, including environmental disasters (Fukuda et al., 1998), pollution
(Mocarelli et al., 2000; Weisskopf et al., 2003), and mothers of advanced age (Orvos et al., 2001).
In the current analysis, we assess indicators of stress and blastocyst quality as a result of BPA
exposure during in vitro bovine oocyte maturation.
70
BPA is an EDC best known for its ability to mimic estrogen, though it has been cited to
have many effects in various physiological systems. The reproductive effects of BPA published
to date are vast. Altered hormone secretion (Vandenberg et al., 2009), decreased implantation
success (Berger et al., 2010; Ehrlick et al., 2012a), altered ovarian morphology (Suzuki et al.,
2002), abnormal early embryonic development (Takai et al., 2001) and recurrent miscarriage
(Kwintkiewicz et al., 2010) are some of the many observed reproductive effects of BPA exposure.
BPA has been measured at about 2.4 ng/mL in follicular fluid of women undergoing IVF (Ikezuki
et al., 2002), and studies examining the effects of BPA at the oocyte maturation phase have found
a variety of effects including delayed cell cycle progression (Can et al., 2005), spindle aberrations
(Can et al., 2005; Eichenlaub-Ritter et al., 2008), misalignment of chromosomes (Hunt et al., 2003;
Eichenlaub-Ritter et al., 2008), centrosomal alterations (Can et al., 2005) and increased aneuploidy
(Hunt et al., 2003; Susiarjo et al., 2007).
Despite the many reported effects of BPA on female fertility and reproduction, the possible
effects of BPA on preimplantation development in mammals are not as well documented. There is
evidence that gestational or perinatal exposure can result in short- and long-term effects with the
possibility of a grand-maternal effect. Developmental exposure to BPA has resulted in poor
pregnancy outcomes such as pre-term birth (Cantonwine et al., 2010), predisposition to the
development of metabolic syndrome (Wei et al., 2011) as well as alterations in gene expression
and behaviour of offspring that may be passed on to the subsequent generation (Wolstenholme et
al., 2012). However, fewer studies have analyzed the effects of early exposure to BPA on
blastocyst development and quality in mammals. Existing studies have found detrimental
outcomes as a result of preimplantation BPA exposure with evidence suggesting that embryo
71
development may be altered as a result of BPA exposure during the preimplantation period
(Tsutsui et al., 1998; Takai et al., 2000, 2001; Xiao et al., 2011; Yan et al., 2013).
As reported in Chapter 1, exposure of bovine oocytes to 30 ng/mL during in vitro
maturation results in decreased maturation success, and increased incidence of MII abnormalities
including a smaller, compressed spindle, and misalignment of the chromosomes at the metaphase
plate. These parameters have been linked to poor developmental competency of the oocyte and
resulting embryos (Rama Raju et al., 2007; Ye et al., 2007; Tomari et al., 2011). Thus, the current
study was designed to evaluate the effects of exposure to BPA during oocyte maturation in vitro
on early embryo development and blastocyst quality. Cleavage and blastocyst rates, sex ratio of
embryos, total cell number, and apoptosis are examined as indicators of embryo quality as well as
stress experienced by the embryo during development. Based on findings in the literature, we
hypothesized that BPA exposure during oocyte maturation would result in decreased
developmental rates, a skewed sex ratio, lower total cell numbers, and a higher proportion of
apoptotic cells, thereby indicating decreased blastocyst quality.
72
MATERIALS AND METHODS
Experimental design
Oocytes were matured in vitro in one of five treatment groups as described in Chapter 1.
Chemicals
All chemicals were obtained from Sigma Life Sciences, Oakville, ON unless otherwise
stated.
Oocyte collection and in vitro embryo production
Procedures for collection and maturation of bovine cumulus-oocyte complexes (COCs)
were performed as described in Chapter 1.
Frozen-thawed Bos taurus semen (EastGen, Guelph, Ontario, Canada) with known in vitro
fertility was used for IVF. Semen was prepared by swim-up in 1.5 mL of sperm-HEPES TALP for
45 minutes at 38.5°C in 5% CO2 in air. Subsequently, swim up product was centrifuged at 2000
rpm for 7 min to obtain a pellet. Mature COCs were washed twice in HEPES/Sperm TALP, twice
in fertilization medium IVF-TALP supplemented with 20 μg/mL heparin, and transferred to 80 μL
droplets containing IVF-TALP under silicone oil (Paisley Products, Scarborough, ON, Canada).
The mature oocytes were co-incubated with a final concentration of 1 x 106 of motile sperm for 20
hours at 38.5°C in 5% CO2 in air.
Subsequently, presumptive zygotes were collected and transferred into a 15 mL conical
tube containing 2 mL sperm-HEPES TALP. Zygotes were vortexed for 120 seconds to remove the
cumulus cells. Stripped zygotes were washed twice in sperm-HEPES TALP and twice in synthetic
oviductal fluid (SOF) media supplemented with sodium pyruvate, non-essential and essential
amino acids, gentamicin, 15% BSA in SOF, and CanSera. Zygotes were then transferred to 30 μL
73
droplets of SOF media under silicone oil (Paisley Products, Scarborough, ON, Canada) and
incubated at 38.5°C and 5% O2.
Calculation of development rates
Cleavage rates were calculated at 48 hours post fertilization by comparing the number of
zygotes that had cleaved to the number of zygotes originally placed into culture. Blastocyst rates
were calculated at 8 days post fertilization by comparing the number of blastocysts that had
developed to the number of zygotes originally placed into culture.
Embryo sexing by PCR
A total number of 233 blastocysts were collected on Day 8 post fertilization and sexing
analysis was carried out as described by Hamilton et al. (2012). Blastocysts were removed from
culture and transferred into 0.2% pronase solution (Sigma-Aldrich, St. Louis, Missouri) for 1-2
minutes to dissolve the zona pellucidae in order to avoid sperm residue between the zona pellucida
and oocyte membrane. Blastocysts were then washed three times in PBS with 0.1% PVA, and
transferred individually to 0.2 mL PCR tubes with minimal PBS-PVA, submersed in liquid
nitrogen and stored at -80°C.
Frozen embryos were thawed and lysed with a lysis solution containing proteinase K using
an MJ Research PTC-200 Thermal Cycler (Bio-Rad Laboratories, Hercules, CA, USA) for one
hour at 37°C, and 15 minutes at 95°C, holding at 4°C at the end. The lysis product was then split
into two, half of which was used to amplify testis specific protein Y-encode (TSPY) and the other
half was used to amplify glyceraldehyde 3-phosphate dehydrogenase (GAPDH) as a reference
gene. Primers used in the sexing analysis are displayed in Table 1. The amplifications were
performed using the following program by real time polymerase chain reaction (qPCR):
74
denaturation at 95°C for 10 seconds, annealing at 65°C for 10 seconds, elongation at 72°C for 10
seconds, and acquisition of fluorescence for 10 seconds.
Table 1 – qPCR primers for sexing analysis
Gene
GAPDH
Genbank accession
number
NM_001034034.1
Primer sequence (5’-3’)
Source
5’-ttcctggtacgacaatgaatttg-3’
Hamilton et
al., 2011
Product
size (bp)
153
5’-ggagatggggcaggactc-3’
TSPY
X74028
Caudle, 2013
- MSc thesis
5’-tgcttcgaggaagacatcg-3’
210
5’-cctcctctgatggttcttgc-3’
TUNEL analysis of apoptosis in blastocysts
Apoptosis in all treatment groups was assessed with the use of terminal deoxynucleotidyl
transferase mediated dUTP nick-end labelling (TUNEL) assay (Roche Diagnostics, Indianapolis,
IN, USA) as previously described (Matwee et al., 2000; Ashkar et al., 2010b). Blastocysts were
collected on day 8 post fertilization and washed three times in PBS containing 0.1% PVA and
fixed for one hour in 4% paraformaldehyde. Fixed blastocysts were stored in 1%
paraformaldehyde at 4°C for a period no longer than two weeks. Subsequently, fixed blastocysts
were permeabilized in 0.5% Triton X-100 supplemented with 0.5% sodium citrate for 1 hour.
Positive control embryos were incubated for 1 hour in a humidity chamber at 38.5°C in a DNase
solution containing 6.5 μL DNase buffer and 5.0 μL DNase I (400 IU/mL; ZyGEM, New Zealand).
Embryos were exposed to fluorescein isothiocyanate (FITC)-conjugated dUTP and
terminal deoxynucleotidyl transferase (TdT) enzyme in a 9:1 ratio for 1 hour in a humidity
75
chamber at 38.5°C. Negative control embryos randomly selected from each treatment group were
incubated with only FITC. Examples of a test embryo, positive control, and negative control are
shown in Figure 8. Following incubation, all groups were treated with 50 μg/mL RNase (Roche
Diagnostics, Indianapolis, IN, USA) for 1 hour in a humidity chamber at 38.5°C. The embryos
were counterstained with 40 μg/mL propidium iodide (PI) for 45 minutes in a humidity chamber
at 38.5°C. Embryos were washed three times and placed onto slides individually. VECTASHIELD
anti-fade solution (Vector Laboratories, California) was placed on top of each embryo. Blastocysts
with TUNEL and PI staining were visualized using an Olympus FV1200 Confocal Microscope
with laser wavelengths of 488 nm for FITC and 543 nm for PI at 20x magnification. Z-stacks were
taken for each embryo at a depth of 1.0 – 5.0 μM. A total of 35, 20, 28, 33, and 32 embryos were
analyzed in the IVM, 0.1% ethanol, 2 μg/mL E2, 15 ng/mL BPA, and 30 ng/mL BPA treatment
groups, respectively.
Cell number and apoptosis rates were analyzed using ImageJ software. Image sequences
were imported and the Cell Counter plugin was used to count blastocyst nuclei by hand. Since
different cell pathways share common biochemical features (Darzynkiewicz et al., 2001; Leist &
Jaattela, 2001), only nuclei displaying both biochemical and morphological features of apoptosis
were classified as apoptotic, as previously described (Favetta et al., 2004) thereby providing a
conservative estimate of apoptosis. This method of categorization of nuclei was adapted from
Gjørret et al. (2003) and nuclei were assigned to one of four categories as follows: (1) Normal:
non-condensed and TUNEL-negative; (2) Condensed: condensed and TUNEL-negative; (3) DNAdamaged: non-condensed and TUNEL-positive; (4) Apoptotic: condensed and TUNEL-positive.
Total cell number was calculated as the sum of all four cell types within a single blastocyst.
76
Condensed, DNA-damaged, and apoptotic nuclei rates were calculated as the number of cells
categorized as such divided by the total cell number of the embryo.
Figure 8. Representative images of test samples (S), and positive (+) and negative (-)
controls. Scale bar = 50 μM.
77
Statistical analyses
Two-tailed Fisher’s exact test was used to analyze the differences in cleavage and
blastocyst formation rate by comparing the proportion of embryos to cleave or reach blast versus
those that did not. The impact of the treatment group on the resulting sex ratio was analyzed by
linear regression analysis, and the correlation between blastocyst rates and sex ratios were
completed using simple linear regression (Graphpad Prism 6). Furthermore, odds ratios were
performed to determine if the sex ratios in each treatment group differed significantly from the
expected 1:1 ratio (MedCalc statistical software). A p value of <0.05 was used to establish
statistical significance.
Total cell number, apoptosis rates, as well as rates of other classified nuclei were
interpreted using Graphpad Prism 6 software. Differences were compared between treatment
groups with one-way ANOVA, and a p value of <0.05 was used to establish statistical significance.
When significance was indicated, Tukey’s multiple comparison test was used to determine
differences between individual treatment groups.
78
RESULTS
Developmental rates, sex ratio, and total cell number of blastocysts
Cleavage and blastocyst rates, sex ratio and total cell number of blastocysts are presented
in Table 1. The cleavage rate of the 30 μg/mL BPA treatment group (64.4%) was significantly
lower compared to both the no-treatment (IVM) and vehicle controls (0.1% ethanol) (79.7%
(p=0.0036) and 77.6% (p=0.0095), respectively). The cleavage rates of the two BPA treatment
groups also differed, with a significantly lower proportion of zygotes cleaving following exposure
to 30 ng/mL during oocyte maturation compared to 15 ng/mL (64.4% vs. 78%, respectively;
p=0.0024). Cleavage rates of the 30 ng/mL BPA treatment group was also lower than the E2 group,
however these results were not statistically significant (64.4% vs. 73%, respectively; p=0.0587).
The proportion of cultured embryos to reach blastocyst was significantly lower in the 30 ng/mL
BPA-exposed group (21.4%) compared to the IVM and 0.1% ethanol groups (36.9% (p=0.0005)
and 29.7% (p=0.0077), respectively). Blastocyst rates in the lower dose of BPA (24.1%) was
significantly lower than that of the IVM group (36.9%, p=0.0381), but not the 0.1% ethanol group
(29.7%) (p=0.2236).
The sex ratio of single blastocysts were calculated for all groups and presented as the
proportion of blastocysts tested that were identified as male (Table 1). The proportion of male
blastocysts arising from oocytes in both BPA groups were lower than controls, with the 30 ng/mL
BPA group exhibiting a significant decrease in the male: female sex ratio compared to the expected
1:1 ratio (p=0.0326), and the 15 ng/mL BPA group showing a non-significant decrease (p=0.0882).
Furthermore, all groups were subject to linear regression analysis to determine the likelihood that
the changes in sex ratio were a result of the treatment administered (p=0.0103). A strong but nonsignificant correlation was also found between the resulting blastocyst rates and sex ratios in each
79
treatment group (p=0.053). Total cell number of blastocysts were calculated and are displayed in
Table 1. The mean blastocyst cell number did not differ between any of the treatment groups and
ranged from 114.3 ± 9.5 in the 30 ng/mL BPA group to 131.7 ± 7.0 in the E2 group (p=0.67).
Nuclear condensation, DNA damage, and apoptosis
Blastocyst nuclei were characterized by their morphological and biochemical
characteristics through TUNEL analysis. Nuclei that were TUNEL-negative, but morphologically
condensed were characterized as condensed nuclei. Nuclei staining positive for TUNEL,
exhibiting the biochemical characteristics of apoptosis, but not condensed were characterized as
DNA-damaged. Finally nuclei exhibiting both the biochemical (TUNEL-positive stain) and
morphological (condensation and/or fragmentation) characteristics of apoptosis were categorized
as apoptotic. Representative images of condensed, DNA-damaged, and apoptotic nuclei are
exhibited in Figures 9, 10, and 11 respectively.
The mean proportion of condensed nuclei did not differ between the treatment groups.
Examples of condensed nuclei that were not apoptotic as well as the mean proportion of condensed
nuclei per blastocyst in all treatment groups are displayed in Figure 9. Representative images as
well as the mean proportion of DNA-damaged nuclei are displayed in Figure 10. Blastocysts that
developed from oocytes exposed to 30 ng/mL BPA during maturation contained a significantly
higher proportion of DNA-damaged nuclei (0.7%) than all of the other groups (0.06% – 0.22%;
p<0.05), though incidence was rare in all treatment groups.
Apoptotic nuclei were stained positive for TUNEL and exhibited morphological
characteristics of apoptosis. Example images as well as the mean proportion of apoptotic nuclei
in blastocysts are displayed in Figure 11. Exposure to 30 ng/mL BPA during oocyte maturation
80
resulted in blastocysts with a significantly higher proportion of apoptotic cells (13.6%) in
comparison to all of the other groups (7 – 9.3% p<0.01).
81
Table 2 – Effect of IVM treatments on development, sex ratio, and total cell number of bovine embryos
Zygotes
Cleaved
Treatment
Day 8 blastocysts
207
Percentage of
zygotes
79.7a
94
Percentage of
zygotes
36.9a
295
235
77.6a
92
259
201
73.0ab
n
n
IVM
255
0.1% ethanol
2 μg/mL E2
Sex Ratio
Blastocyst cell number
n
Mean ± SEM
54
Percentage
Male
49.9
35
124.1 ± 8.3
29.7ab
40
54.1
20
118.7 ± 13.6
79
27.4abc
43
53.8
28
131.7 ± 7.0
a
91
24.1
bc
50
38.4
33
121.4 ± 7.1
81
21.4c
46
34.1*
32
114.3 ± 9.5
15 ng/mL BPA
321
260
78.0
30 ng/mL BPA
323
228
64.4b
n
abc
n
Values in the same column with different superscripts are significantly different (p<0.05).
(*) indicates significant variation in sex ratio from the expected 1:1 ratio; *p<0.05
82
Figure 9. Condensed nuclei (white arrows) in blastocysts. Representative images (A) and
mean proportions (B) of condensed nuclei without TUNEL staining in blastocysts in each of the
treatment groups. Mean ± SEM. Scale bars = 50 μM.
83
Figure 10. DNA-damaged nuclei (white arrow) in blastocysts. Representative images (A)
and mean proportions (B) of TUNEL-positive, non-condensed nuclei in blastocysts in each of
the treatment groups. Mean ± SEM. Analysis of variance (ANOVA) and Tukey’s multiple
comparison test, *p<0.05. Scale bars = 50 μM.
84
Figure 11. Apoptotic nuclei (white arrows) in blastocysts. Representative images (A) and
mean proportions (B) of TUNEL-positive, condensed nuclei in blastocysts in each of the
treatment groups. Mean ± SEM. Analysis of variance (ANOVA) and Tukey’s multiple
comparison test, **p<0.01. Scale bars = 50 μM.
85
DISCUSSION
The results presented in this study indicate that exposure of bovine oocytes to 30 ng/mL
BPA during in vitro oocyte maturation decreases embryonic cleavage and blastocyst rates, skews
sex ratio towards femaleness, and increases apoptosis, as well as the proportion of non-apoptotic
cells undergoing DNA damage, in blastocysts. However, the total cell number of blastocysts did
not differ between any of the treatment groups. The lower dose of BPA (15 ng/mL), resulted in a
decrease in blastocyst rate, but only in comparison to the no-treatment control, and a skewed sex
ratio in comparison to the expected 1:1 ratio, but this did not reach statistical significance. No other
indicators of blastocyst quality were altered in the 15 ng/mL BPA group. Taken together these
findings suggest that oocyte exposure to low levels of BPA during oocyte maturation can decrease
developmental potential and quality of the resulting blastocyst. Other studies have suggested that
BPA has the potential to affect early embryo development, however results between studies are
variable (Tsutsui et al., 1998; Takai et al 2000, 2001; Bloom et al., 2011b; Xiao et al., 2011; Ehrlich
et al., 2012b; Yan et al., 2013).
In vitro studies examining BPA exposure during preimplantation development have
exhibited similar results, and the same variability, as in vivo studies. Takai et al. (2000, 2001)
observed opposite effects of low (1-3 nM; <1 ng/mL) and high (100 μM; 22.83 μg/mL) doses of
BPA on murine embryos. Two-cell embryos exposed to 100 μM resulted in fewer embryos
reaching the blastocyst stage, whereas treatment with 1nM BPA resulted in a greater number of
embryos reaching blast. Blastocyst morphology, total cell number, and sex ratio did not differ
between treatment groups and the control (Takai et al., 2000). Both the high and low treatments
led to pups that were significantly heavier than controls at weaning (Takai et al., 2001). Thus BPA
exposure during the preimplantation period has the potential to affect blastocyst development, as
86
well as induce postnatal effects. However this is the first study, to our knowledge, to investigate
the effects of BPA exposure during in vitro maturation of oocytes on embryo development and
blastocyst quality in the bovine species.
Previously we have shown that BPA exposure during oocyte maturation under the same
conditions as were used currently, results in only a small proportion of BPA being taken up by the
oocyte (Chapter 1). Oocytes exposed to 30 ng/mL during maturation resulted in an average uptake
level of 2.48 ng/mL per oocyte, whereas the lower dose, 15 ng/mL resulted in an oocyte
concentration of 1.69 ng/mL. Since the amount of BPA taken up by an oocyte is likely of greater
importance than the initial exposure dose, it is essential to consider uptake of BPA when assessing
developmental effects. Additionally, we observed that maturational exposure decreased the
success of oocytes reaching MII, and an increased incidence of MII spindle abnormalities
including altered spindle morphology and misalignment of the chromosomes at the metaphase
plate (Chapter 1). Thus the effects observed in this study are likely the result of alterations made
during oocyte maturation, and differ from similar studies in which exposure occurs during embryo
development.
Embryo development, sex ratio, and blastocyst cell number
As mentioned, blastocyst development was delayed as a result of embryonic exposure to
100 μM BPA in vitro (Takai et al., 2000), and urinary BPA concentrations have been associated
with decreased embryo development (Ehrlich et al., 2012b). The results presented here suggests
that exposure to BPA during oocyte maturation decreases the embryo’s ability to cleave following
fertilization, and develop to the blastocyst stage. Total cell number and sex of embryos that did
reach blastocyst were determined to further analyze blastocyst quality. The sex ratio was
significantly affected by oocyte maturational exposure to 30 ng/mL BPA, with a 15.8 - 20%
87
decrease in sex ratio, meaning 34.1% of blastocysts in the 30 ng/mL group were male compared
to 49.9% in the IVM group and 54.1% in the 0.1% ethanol group. We found no differences in total
cell number of blastocysts between all of the treatment groups.
Sex ratio was analyzed as it is considered to be an indicator of embryo quality and can give
an idea of the severity of stress experienced by the embryo. For instance, sex ratio has been altered
as a result of maternal stress in rodents (Lane & Hyde, 1973; Moriya et al., 1978; Pratt & Lisk,
1989; Krackow, 1997; James, 1998) and humans (Fukuda et al., 1998; Mocarelli et al., 2000; Orvos
et al., 2001; Weisskopf et al., 2003). In vitro, blastocyst sex ratio has been found in many instances
to be altered by environmental conditions such as hyperglycemia, oxidative stress, in vitro culture
media composition, and exposure to EDCs (Mocarelli et al., 2000; Weisskopf et al., 2003
Gutiérrez-Adán et al 2001a; Larson et al., 2001; Ishihara et al., 2007). Increased glucose levels in
in vitro culture media appears to favour male embryos and inhibit female embryo development
(Gutiérrez-Adán et al 2001a; Larson et al., 2001), whereas paternal exposure to the dioxin TCDD
resulted in skewed sex ratio in favour of female embryos (Ishihara et al., 2007).
Studies from our lab have previously indicated a skew in sex ratio as a result of
environmental conditions. In vitro maturation of bovine oocytes in an open-well glassware system
with ethanol-supplemented media resulted in a sex skew in favour of females (34% males)
(Macaulay et al., 2011). This skew in sex ratio was accompanied by a decrease in cleavage and
blastocyst rates versus the standard control IVM system, indicating that alterations in IVM
protocols and culture environment of oocytes during maturation can have developmental
consequences such as decreased developmental potential and embryo quality (Macaulay et al.,
2011), which is in line with the results currently obtained. Furthermore, our lab has previously
described a positive correlation between blastocyst formation rates and the percentage of male
88
embryos, indicating that a decrease in blastocyst rates was accompanied by a decrease in the
proportion of male embryos (Macaulay et al., 2013).
These trends indicate that male embryos are not as resilient as female embryos and may be
less likely to survive under stressful conditions and support the Trivers and Willard hypothesis that
postulates that an excess of male offspring are favoured by natural selection when conditions, and
likelihood to survive, are good, and mothers in poor conditions benefit most by the production of
daughters (Trivers & Willard, 1973). This may be a result of differential survival rates of male and
female embryos during preimplantation development. Similar to the above-mentioned studies, the
male:female sex ratio in the current analysis was decreased, meaning there was a higher proportion
of female blastocysts following exposure to 30 ng/mL BPA during oocyte maturation than the
control groups. The skew in sex ratio may indicate embryonic stress, to which male embryos are
less likely to survive than female embryos. Determination of the sex of arrested embryos would
provide valuable information on whether male embryos are arresting at a higher rate than female
embryos.
Nuclear condensation, apoptosis, and DNA-damage
Apoptosis in the blastocyst is common, and is an important mechanism by which the
embryo can remove damaged cells or cells with abnormal developmental potential without
affecting neighbouring cells (Jacobson et al., 1997; Betts & King, 2001; Hansen & Fear, 2011).
An increased incidence of apoptosis may indicate an embryonic response to stress. The ability of
the embryo to induce apoptosis in times of stress may help it to survive suboptimal conditions,
however the extent to which apoptosis occurs in the blastocyst plays an important role in the fate
of that embryo (Byrne et al 1999; Betts & King, 2001; Paula-Lopes & Hansen, 2002b; Hansen &
Fear, 2011).
89
In the current study, exposure of oocytes during maturation to 30 ng/mL BPA resulted in
a significantly higher proportion of cells undergoing apoptosis at the blastocyst stage. BPA has
previously been found to increase apoptosis in embryonic midbrain cells in vitro (Liu et al., 2013),
and murine antral follicles following maternal exposure to BPA (Peretz et al., 2013). Furthermore,
apoptosis in the blastocyst has been induced by stressors such as heat stress and exposure to
xenobiotic chemicals (Betts & King, 2001; Hamdoun & Epel, 2007). The increased incidence of
apoptosis currently observed may indicate a decreased developmental potential, possibly caused
by embryonic stress and/or cellular abnormalities as a result of exposure to BPA during oocyte
maturation. Further analysis is required to assess the pathway through which apoptosis is being
induced by BPA exposure.
Morphologically, apoptotic cell death differs from other forms of cell death, such as
necrosis, by the presence of chromatin condensation and nuclear fragmentation (Majno & Joris,
1995). It is important to distinguish between nuclei which display both the biochemical and
morphological characteristics of apoptosis, from those which display one or the other (Gjørret et
al., 2003). Observation of more than one of the characteristics of apoptosis is essential for proper
identification (Darzynkiewicz et al., 2001). Nuclei displaying morphological characteristics of
apoptosis are commonly observed in embryos after the 8-cell stage (Gjørret et al., 2003). Nuclear
condensation may be a result of early apoptosis in which DNA fragmentation is yet to occur
(Collins et al., 1997), or a misinterpretation of prophase nuclei due to the higher chromatin content
than interphase nuclei (Gjørret et al., 2003). Alternatively, nuclei may condense and become
fragmented by alternative mechanisms such as abnormal mitotic chromosome segregation, which
can result in micronuclei-like structures (Gjørret et al., 2003). Currently, the proportion of nonapoptotic condensed nuclei did not differ between treatment groups.
90
TUNEL allows for detection of apoptotic cells in situ by labelling DNA fragmentation.
However, DNA fragmentation is not exclusive to apoptotic cell death and all nuclei undergoing
DNA fragmentation, not just apoptotic nuclei, will stain positive for TUNEL. Thus it is important
to distinguish apoptotic cells from non-apoptotic cells that contain DNA fragmentation. Nuclei
displaying the biochemical features of apoptosis in absence of morphological features were
therefore not considered to be apoptotic. The proportion of cells displaying normal morphology
with DNA damage (TUNEL-positive) presented here was significantly higher in the 30 ng/mL
BPA group than all of the other groups. Therefore in addition to the increased apoptosis rate in the
30 ng/mL BPA group, there was also evidence of increased non-apoptotic DNA fragmentation.
Further analysis is required to determine the cause of this DNA damage and whether it represents
an alternative form of cell death.
Taken together, the findings presented indicate that exposure to low levels of BPA during
in vitro oocyte maturation has the potential to result in decreased embryonic developmental
potential and quality as evidenced by decreased development rates, skewed sex-ratio, and
increased proportion of cells exhibiting apoptosis and non-apoptotic DNA damage. The maturing
oocyte is vulnerable to environmental perturbations and can influence further development of the
embryo. In Chapter 1, we observed that the same experimental conditions resulted in low levels of
BPA taken up by the oocyte. Exposure to 15 ng/mL BPA resulted in an average uptake of 1.69
ng/mL and exposure to 30 ng/mL resulted in an average uptake of 2.48 ng/mL BPA. As we have
previously observed meiotic aberrations under the same conditions (Chapter 1), it is possible that
irregularities resulting from an abnormal spindle and chromosome misalignment could result in
aneuploid blastomeres, which could be removed from the blastocyst via apoptosis. There are
various causes of stress in the preimplantation embryos and these stressors can result in increased
91
apoptosis and/or a skewed sex ratio. In some cases activation of stress pathways, as has been
observed in heat and oxidative stress, may induce apoptosis by altering gene expression in the
preimplantation embryo (Kawarsky & King, 2001; Camargo et al., 2007; Rho et al., 2007).
Mechanistic analyses are essential to determine how BPA exposure during oocyte maturation is
disrupting the development and quality of the early embryo.
92
CHAPTER THREE
Gene expression of developmentally important genes in Bos taurus MII oocytes and
blastocysts following bisphenol A exposure during in vitro oocyte maturation.
93
INTRODUCTION
Integrity of early embryonic development is influenced by the environmental conditions
surrounding the embryos or the oocyte from which it developed. Alterations in the oocyte’s
microenvironment can influence not only oocyte metabolism and developmental competence, but
also embryo quality and gene expression patterns (Guérin et al., 2001; Leroy et al., 2012).
Environmental conditions, such as in vitro culture conditions, temperature change, oxygen tension
change, and toxic exposures can cause abnormalities in oocyte maturation, embryonic
development, lead to poor developmental outcomes, and alter gene expression of the early embryo
(Niemann & Wrenzycki, 2000; Li et al., 2005; Lonergan et al., 2006; Camargo et al., 2008).
Furthermore, changes in gene expression of the mature oocyte may be indicative of a disruption
to developmental programs (Paczowski et al., 2011; Yuan et al., 2011).
Although many aspects of oocyte maturation remain unknown, it has been well established
that a complex variety of factors are involved in meiotic progression, and spindle assembly,
integrity, and localization. Meiosis is largely regulated by the cell cycle regulator maturationpromoting factor (MPF). MPF is one of the main signalling pathways of oocyte maturation and
one of the key regulators of cell cycle progression and oocyte maturation (reviewed by Voronina
& Wessel, 2003). It consists of two subunits, the catalytic cyclin-dependent kinase 1 (CDK1) (aka
CDC2, p34cdc2), an important cell cycle regulator, and the regulatory cyclin B (Dunphy et al., 1988;
Gautier et al., 1988, Pines et al., 1989; Gautier et al., 1990). Additionally, genes important to
spindle assembly, function, and/or translational regulation include aurora kinase A (AURKA)
(Uzbekova et al., 2008), deleted in azoospermia-like (DAZL) (Chen et al., 2011), and kinesin
family member 5b (KIF5b) (Brevini et al., 2007; Kidane et al., 2013). KIF5b is a motor molecular
and is critical to germinal vesicle breakdown (GVBD), kinetochore assembly, chromosome
94
stability, polar body extrusion, integrity and polarity of the mitotic spindle, as well as metaphase
alignment (Kidane et al., 2013). AURKA has been found to be essential for centrosome and spindle
assembly, chromosome attachment, and chromosome alignment (Marumoto et al., 2005;
Uzbekova et al., 2008). DAZL is a microtubule regulator and directs translation of various genes
important to cell cycle regulation, chromatin remodeling, spindle function, and centrosome and
spindle assembly (Chen et al., 2011). Alterations in these genes during oocyte maturation has
resulted in impaired embryonic development (KIF5b) (Tanaka et al., 1998; Takamiya et al., 2004),
meiotic arrest and increased aneuploidy in oocytes (AURKA) (Uzbekova et al., 2008; Lane et al.,
2010), and disruption to meiotic spindle assembly (DAZL) (Chen et al., 2011).
Alterations in expression of stress- and metabolism- related genes in the MII stage may
leave the resulting embryo unequipped for early embryonic development (Brevini et al., 2002;
Percell & Moley, 2009). In the embryo, stress can result in alterations of stress-responsive genes
including heat shock protein 70 (HSP70), tumour protein p53, and glucose transporter 1 (GLUT1)
(Batt et al., 1991; Kawarsky & King, 2001; Camargo et al., 2007; Rho et al., 2007; Hu et al., 2011).
Expression levels of these genes have been shown to vary as a result of embryonic stress, and these
responses appear to depend on both the level of stress experienced and the stage of development
at which the stress occurs (Wrenzycki et al., 1998; Niemann & Wrenzycki, 2000; Kawarsky &
King, 2001; Peretz et al., 2012). In addition to mRNA expression alterations as a result of stress
experienced during embryonic development, the environment in which the oocyte matures has also
been found to influence gene expression in the mature oocyte and the resulting embryo (Russel et
al., 2006; Sagirkaya et al., 2007; Wells & Patrizio, 2008; reviewed by Virant-Klun et al., 2013).
Among the markers that allow us to evaluate embryonic viability, the expression of
hormone receptors is of significant importance at different stages of development and especially
95
at the oocyte and blastocyst stages (Beker-van Woudenberg et al., 2004; Vasquez & DeMayo,
2013). Alterations in hormone receptor mRNA synthesis may indicate poor developmental
programming or improper conditions, and may be predictive of undesirable reproductive
outcomes. Nuclear receptors play an important role in the oocyte and early embryo (Beker-van
Woudenberg et al., 2004), and are essential for implantation or attachment (reviewed by Vasques
& DeMayo, 2013). The steroid hormone, estradiol (E2) plays a critical role in fertility, oocyte
development, and early embryonic development. E2 primarily exerts its actions by binding to its
receptors ERα and ERβ. Expression of ERs by the embryo may be critical to its survival (Hou &
Gorski, 1993). Thyroid hormones (THs), which activate the TH receptors TRα and TRβ, play a
beneficial role in preimplantation development including increased blastocyst and hatching rates
(Ashkar et al., 2010b, 2013). Both the ERs and TRs have been found to be altered transcriptionally
by exposure to the endocrine disruptor bisphenol A (BPA) (Heimeier et al., 2009; Aghajanova &
Giudice, 2011), but the influence of BPA on mammalian embryo development on ER and TR
expression, as well as that at the oocyte maturation stage, is not fully understood.
In the current study, oocyte and blastocyst gene expression were evaluated under various
treatment conditions occurring solely during oocyte maturation. Genes involved in oocyte
maturation and spindle assembly (CDC2, AURKA, DAZL, KIF5b, alpha tubulin [TUBA]),
response to stress and/or metabolism (HSP70, p53, GLUT1), and hormone receptor genes (ERβ,
TRβ), were quantified in germinal vesicle (GV) stage and metaphase II (MII) oocytes. Blastocysts
resulting from oocytes under the same treatment conditions were further analyzed for variations in
gene expression of the stress, metabolism, and hormone receptor genes. As increased transcript
degradation due to poor environmental conditions may lead to a decrease in mRNA in MII oocytes,
we hypothesized that transcripts in the oocyte would be decreased after BPA treatment during
96
maturation, with the exception of TUBA, which we speculated would not be altered as it has
previously been used as a reference gene in BPA analyses (Gentilcore et al., 2012). Furthermore,
due to existing reports of both embryonic response to stress and BPA action on these genes in
alternate tissues (Niemann & Wrenzycki, 2000; Wrenzycki et al., 2001; Matthews et al., 2001;
Kang et al., 2002; Moriyama et al., 2002; Takao et al., 2003; Levy et al., 2004; Lahnsteiner et al.,
2005; Schirling et al., 2006; Planelló et al., 2008; Heimeier et al., 2009; Vandenberg et al., 2009;
Guo et al., 2012; Liu et al., 2013; Park & Kwak, 2013; Peretz et al., 2013), we have hypothesized
that in the blastocyst, p53, HSP70, and ERβ would be increased as a result of BPA exposure during
oocyte maturation in vitro, whereas GLUT1 and TRβ would be decreased.
97
MATERIALS AND METHODS
Experimental design
Oocytes were matured in vitro in one of five treatment groups as described in Chapter 1.
Chemicals
All chemicals were obtained from Sigma Life Sciences, Oakville, ON unless otherwise
stated.
Oocyte collection and in vitro embryo production
Procedures for oocyte collection and in vitro embryo production are described in Chapters
1 and 2.
RNA extraction
Oocytes from three biological replicates were collected in pools of 40 at GV and at MII
following the various treatments. Embryos from three groups were collected in pools of 5 at the
blastocyst stage. Oocytes were stripped, as described in Chapter 1, and oocytes and blastocysts
were washed three times in PBS-PVA 0.1%, snap-frozen in liquid nitrogen and stored at -80°C.
RNA was isolated from pooled embryos using the AllPrepTM DNA/RNA Micro Kit (QIAGEN,
Inc., Burlington, ON), following the manufacturer’s instructions. In brief, Buffer RLT plus was
added to lyse the cells. Samples were vortexed for homogenization and transferred to an AllPrep
DNA spin column to bind genomic DNA. Flow through was mixed with 70% ethanol and run
through an RNeasy MinElute spin column to bind total RNA. The column was then washed and
dried, followed by RNA elution in RNase-free water. RNA samples were reverse transcribed
immediately following extraction using the one-step protocol with qScriptTM cDNA SuperMix
(Quanta Biosciences, Canada) following the manufacturer’s instructions. cDNA samples were
stored at -20°C until needed.
98
Gene expression analysis
Quantitative real-time PCR (qPCR) was used to measure mRNA expression profiles of
selected genes in the five treatment groups and in the GV oocyte group. Each analysis was
performed on three biological replicates with three technical replicates each. Glyceraldehyde-3phosphate dehydrogenase (GAPDH) and peptidylprolyl isomerase A (PPIA) were used as
reference genes for all qPCR analyses. Relative quantity of target genes were log-transformed and
normalized to relative quantity of the reference genes, GAPDH and PPIA, across samples (ΔΔCq)
(Bio-Rad CFX Manger 3.1). Therefore values presented in figures in the current chapter are
relative normalized gene expression levels.
The primer sequences are listed in Table 3. The expression of all genes were previously
detected in domestic cattle as confirmed by the references provided in Table 3. qPCR was carried
out using the Bio-Rad CFX96 Real-Time PCR System and products were detected with SsoFast
TMEvaGreen® Supermix (Bio-Rad Laboratories, Hercules, CA, USA) according to the
manufacturer’s instructions. Each reaction contained 5 μL of the SsoFastTM EvaGreen®
Supermix reaction mix, 1 μL of a mix of the forward and reverse primers (at a 0.1 μM concentration
for each set of primers, except the ERβ primers, used at a concentration of 0.5 μM) and 2 μL cDNA
for all genes with the exception of ERβ (3uL). The final volume was adjusted to 10 μL using
RNase-free water (Ref Ambion PCR-grade water). A standard curve was established for each
primer set using ovarian tissue cDNA template in five serial dilutions and primers efficiencies
were calculated and used in the analysis. The amplification program was as follows: preincubation
for EvaGreen® Supermix polymerase activation at 95°C for 10 minutes, followed by 50
amplification cycles of denaturation at 95°C for 10 seconds, annealing at 65°C for 10 seconds,
elongation at 72°C for 10 seconds, and acquisition of fluorescence for 10 seconds. After the last
99
cycle, fluorescence acquisition was begun at 72°C, and measurements were taken every 0.5°C until
95°C to generate the melting curve.
Statistical analysis
Expression levels were calculated relative to those of internal reference genes GAPDH and
PPIA in Bio-Rad CFX Manager 3.1. Differences in MII and blastocyst gene expression levels
between treatment groups were analyzed with One-way analysis of variance (ANOVA), and when
statistical significance was observed (p < 0.05), Tukey’s multiple comparison tests were performed
to evaluate differences between specific treatment groups within a single gene (Graphpad Prism
6.0). Differences in expression levels between GV and MII stages were analyzed for each gene
using unpaired student’s t-test (Graphpad Prism 6.0). Differences at p < 0.05 were considered
statistically significant.
100
Table 3. List of primers used for qPCR
Gene
GAPDH
PPIA
p53
HSP70.1
ERβ
TRβ
GLUT1
Genbank accession
number
NM_001034034.1
NM_178320.2
BC102440
U09861
AF110402
281061
NM_174602.2
Source
Hamilton et
al., 2011
N/A
Piper et al.,
2008
Camargo et
al., 2007
Connor et al.,
2005
Ashkar, 2013
(PhD thesis)
Primer sequence (5’-3’)
Primer
efficiency
(%)
102.3
Product
size (bp)
5’-ggagatggggcaggactc-3’
5’-tcttgtccatggcaaatgctg-3’
99.9
111
5’-tttcaccttgccaaagtaccac-3’
5’-ctttgaggtgcgtgtttgtg-3’
100.8
161
5’-gtggtttcttctttggctgtg-3’
5’-aacaagatcaccatcaccaacg-3’
102.8
275
5’-tccttctccgccaacgtgttg-3’
5’-acctgctgaatgctgtgac-3’
103.6
128
5’-gttactggcgtgcctgac-3’
5’-atctatgatcctgccaaacg-3’
104.4
119
5’-tgacacaacacagggaaactg-3’
5’-ctgatcctgggtcgcttcat-3’
98.8
68
5’-ttcctggtacgacaatgaatttg-3’
BermejoÁlvarez et al.,
2010
5’-acgtacatgggcacaaaacca-3’
101
153
TUBA
T90759
CDC2
NM_174016
KIF5B
NM_001192365.1
DAZL
NM_001081725.1
AURKA
DQ334808
Laurent et al., 5’-gctgtggaaacctggg-3’
2000
5’-cattgaccttgtgttggac-3’
Uzbekova et
5’-atggttggatctgctctcg-3’
al., 2008
5’-cattaaagtacggatgattcagtgc-3’
Macaulay et
5’-gcctctaaggaagaagtaaagg-3’
al., 2014
5’-cttgctaaacttgccgattcc-3’
Zhang et al.,
5’-cagcctccaaccatgataaaccc-3’
2008
5’-cataactcctttgttccccagcag-3’
Uzbekova et
5’-tcgggaggacttggtttctt-3’
al., 2008
5’-tgtgcttgtgaaggaacacg-3’
102
103.5
196
103.3
89
100.9
149
115.9
154
92.9
234
RESULTS
mRNA expression in GV and MII oocytes
Expression levels in GV and MII oocytes of the various genes analyzed are shown in Figure
12. The genes studied showed variability in mRNA levels before and after maturation. There were
no significant differences observed between GV and MII stages in ERβ (Figure 12 A), GLUT1
(Figure 12 B), and HSP70 (Figure 12 C) mRNA expression levels. mRNA levels for CDC2 (Figure
12 D), TRβ (Figure 12 E), and TUBA (Figure 12 F) were lower in MII oocytes (p < 0.05), whereas
those for DAZL (Figure 12 G), p53 (Figure 12 H), KIF5b (Figure 12 I), and AURKA (Figure 12
J), were increased in MII oocytes compared to GV (p<0.05), however the increase in KIF5b was
not statistically significant.
Genes involved in oocyte maturation and spindle assembly
The mRNA expression levels of genes involved in meiosis progression, spindle assembly,
integrity, and chromosome stability and alignment following IVM in the various treatment groups
are shown in Figure 13. Significant differences between groups were observed in the CDC2,
AURKA, and DAZL analyses. MII oocytes matured with 15 ng/mL BPA exhibited significantly
higher expression of CDC2 and DAZL mRNA than those matured under standard conditions
(IVM) (Figure 13 A and D, respectively). A similar trend was observed in the KIF5b group,
however these results were not significant. A vehicle effect was observed in several of the genes
studied in the oocyte, though this effect was only significantly different between the no-treatment
and vehicle controls in the AURKA analysis (Figure 13 B). No significant differences were
observed in the TUBA mRNA expression analysis, though there appears to be a non-significant
vehicle effect.
103
Hormone receptor gene expression
Differences in mRNA levels of ERβ (A) and TRβ (B) in MII oocytes following maturation
under various treatments are shown in Figure 14. There were no significant differences in ERβ
mRNA expression in MII oocytes, however changes were observed in TRβ mRNA expression (p
< 0.05). Similar to results observed regarding CDC2 and DAZL mRNA expression, TRβ
expression in MII oocytes following maturational exposure to 15 ng/mL BPA was increased
compared to the no-treatment control (Figure 14 B).
Relative mRNA abundance of ERβ and TRβ in blastocysts resulting from oocyte matured
under various treatments is presented in Figure 15. There were no significant differences in mRNA
expression of ERβ or TRβ between the different treatment groups at the blastocyst stage. Although
there appears to be an elevation of ERβ mRNA expression in the 15 ng/mL BPA group, this did
not reach statistical significance (Figure 15 A).
Genes involved in Stress and Metabolism
The mRNA expression profiles of HSP70 (A), p53 (B), and GLUT1 (C) in MII oocytes are
shown in Figure 16. Significant differences in p53 mRNA expression between treatment groups
were observed (p < 0.05). Expression of p53 mRNA was significantly increased in the 15 ng/mL
BPA group relative to the IVM and the 30 ng/mL BPA groups (Figure 16 B). There were no
significant differences in HSP70 and GLUT1 mRNA expression between treatment group (Figure
16 A and C, respectively). Relative mRNA expression in blastocysts of HSP70, p53, and GLUT1
transcripts is shown in Figure 17. There were no differences in the relative expression levels of
these transcripts at the blastocyst stage between treatment groups.
104
Figure 12. mRNA expression profiles of GV and MII oocytes in ERβ (A), GLUT1 (B), HSP70 (C), CDC2 (D), TRβ (E), TUBA (F),
DAZL (G), p53 (H), KIF5b (I), and AURKA (J). Unpaired t-test; (*) indicates statistical significance, *p<0.05; **p<0.01.
Experiments were conducted on three different pools of 40 oocytes (n = 3) and replicated three times (r = 3) for each sample
105
Figure 13. mRNA expression levels (Mean ± SEM) of CDC2 (A), AURKA (B), KIF5b (C), DAZL (D), and TUBA (E) in MII oocytes
following IVM in respective treatment groups. One-way ANOVA and Tukey’s multiple comparison test. Bars with different letters
indicate statistical significance (p<0.05; **p<0.01). Experiments were conducted on three different pools of 40 oocytes (n = 3) and
replicated three times (r = 3) for each sample.
106
Figure 14. mRNA expression levels of ERβ (A) and TRβ (B) in MII oocytes following IVM in
respective treatment groups. One-way ANOVA and Tukey’s multiple comparison test. Bars with
different letters indicate statistical significance (p<0.05). Experiments were conducted on three
different pools of 40 oocytes (n = 3) and replicated three times (r = 3) for each sample.
107
Figure 15. mRNA expression levels of ERβ (A) and TRβ (B) in blastocysts arising from oocytes
matured in vitro in respective treatment groups. One-way ANOVA and Tukey’s multiple
comparison test. Bars with different letters indicate statistical significance (p<0.05). Experiments
were conducted on three different pools of 40 oocytes (n = 3) and replicated three times (r = 3) for
each sample.
108
Figure 16. mRNA expression levels of HSP70 (A), p53 (B), and GLUT1 (C) in MII oocytes
following IVM in respective treatment groups. One-way ANOVA and Tukey’s multiple
comparison test. Bars with different letters indicate statistical significance (p<0.05). Experiments
were conducted on three different pools of 40 oocytes (n = 3) and replicated three times (r = 3) for
each sample.
109
Figure 17. mRNA expression profiles of HSP70 (A), p53 (B), and GLUT1 (C) in blastocysts
arising from oocytes matured in vitro in respective treatment groups. One-way ANOVA and
Tukey’s multiple comparison test. Bars with different letters indicate statistical significance
(p<0.05). Experiments were conducted on three different pools of 40 oocytes (n = 3) and replicated
three times (r = 3) for each sample.
110
DISCUSSION
The results presented in this chapter showed that there were little to no effects of 30 ng/mL
BPA treatment during oocyte maturation on gene expression in MII oocytes and blastocysts.
Treatment with 15 ng/mL BPA resulted in increased expression of CDC2, AURKA, DAZL, TRβ
and p53 in MII oocytes relative to that of the IVM group. This increased p53 expression resulting
from the 15 ng/mL BPA group was also significantly greater than the expression levels resulting
from 30 ng/mL BPA. There appeared to be a slight vehicle effect, with the vehicle group (0.1%
ethanol) resulting in significantly increased expression of AURKA mRNA and non-significant
increases in several other genes analyzed. There were no significant differences observed in
blastocyst mRNA expression levels of the genes analyzed.
The results currently presented indicate that BPA exposure during oocyte maturation may
alter transcript levels of developmentally important genes in MII oocytes, and the nature of these
alterations is dictated by exposure dose. A number of the genes analyzed showed a trend exhibiting
decreased mRNA expression in MII oocytes matured with 30 ng/mL BPA compared to the vehicle
control, however these results were not statistically significant and require further investigation to
reach definitive conclusions. As mentioned, a vehicle effect was observed and increased
expression as a result of 15 ng/mL BPA was partly due to the ethanol vehicle. Increased mRNA
expression as a result of ethanol exposure has previously been observed in bovine MII oocytes
(Gomez et al., 2004) and porcine COCs (Lee et al., 2014a). The concentrations used were much
higher in these studies than that used in the current study, and we did not observe developmental
effects similar to those that were associated with the higher doses (1% and 3%) (Lee et al., 2014a).
Since we did not observe developmental effects as a result of ethanol exposure in Chapters 1 and
2, the changes observed as a result of ethanol exposure do not appear to be functionally significant,
111
at least up to the blastocyst stage, however they must be considered when using ethanol in media
for oocyte maturation.
mRNA expression in GV and MII oocytes
In the current study, we analyzed changes in mRNA expression between the GV and MII
stages to offer perspective of the results obtained (Figure 12). Of the genes analyzed 3 were
significantly downregulated in MII oocytes (CDC2, TRβ, and TUBA), and 4 genes (DAZL, p53,
KIF5b, and AURKA) were upregulated in MII oocytes however expression levels in KIF5b were
not significantly different. Results currently observed in GV and MII oocyte expression of KIF5b,
CDC2, p53, HSP70, and AURKA are in line with previous reports (Manejwala et al., 1991;
Juriscova et al., 1998; Swain et al., 2008; Uzbekova et al., 2008; Mamo et al., 2011; Niu et al.,
2013; Macaulay et al., 2014). There was no observable correlation between GV/MII pattern and
gene-specific BPA effect.
Meiosis and spindle assembly
Due to the ability of BPA to induce meiotic aberrations during oocyte maturation as was
observed in Chapter 1, we analyzed the expression of several meiosis and spindle-related genes in
MII oocytes. Significant differences were observed between treatment groups of three of the genes
studied, (CDC2, AURKA, and DAZL) (Figure 13). CDC2, AURKA, and DAZL expression in
MII oocytes was significantly increased in MII oocytes following exposure to 15 ng/mL BPA
relative to that of the IVM group. Additionally, expression profiles for CDC2, AURKA, and KIF5b
showed similar patterns of expression between the treatment groups, characterized by nonsignificant decreases in the 30 ng/mL BPA and E2 groups compared to the vehicle control, with
no change or an increase in expression in the 15 ng/mL BPA groups. Further investigation is
required to determine if the higher dose of BPA is interfering with processing of these genes that
112
may not be apparent at the mRNA level. Differences not observed through the current analysis,
such as protein levels or timing of gene expression changes, will further help to elucidate the
functional significance of the current findings and how BPA exposure is resulting in the meiotic
defects observed under the same conditions in Chapter 1.
BPA exposure has previously resulted in upregulation of CDC2 expression in amalone
embryos (Zhou et al., 2011), DAZL expression in mouse embryonic stem cells undergoing
differentiation (Aoki & Takada, 2012), and AURKA expression in the human breast epithelial cell
line MCF-10F (Fernandez et al., 2012). Conversely, DAZL and AURKA expression levels were
decreased following BPA exposure in fetal germ cells (Zhang et al., 2012) and fetal mouse ovary,
respectively (Lawson et al., 2012). These genes, as well as KIF5b which has not been studied
following BPA exposure, are important to various aspects of the cell cycle and spindle assembly,
and are thus critical regulators of meiosis (Andresson et al., 1998; Mendez et al., 2000; Yao et al.,
2004; Uzbekova et al., 2008; Baumann et al., 2010; Chen et al., 2011). Alterations in mRNA
expression of these genes during maturation has previously resulted in developmental arrest
(CDC2) (reviewed by Uzbekova et al., 2008), disruptions to spindle assembly (DAZL) (Chen et
al., 2011), and increased incidence of aneuploidy and chromosome alignment (AURKA) (Swain
et al., 2008; Shuda et al., 2009; Lane et al., 2010).
Thus given BPA’s ability to disrupt meiosis and spindle formation (Can et al., 2005; Lenie
et al., 2008; Machtinger et al., 2013; Ferris et al., 2015, Chapter 1), investigation of genes important
to spindle assembly and function during meiosis is vital to analyze how these spindle effects may
result. Further analysis observing polyadenylation, translation, and degradation of these genes is
required to more fully analyze BPA’s implications regarding these genes during oocyte maturation.
Furthermore, due to recent evidence that the cumulus cells can transport RNA molecules into the
113
oocyte during maturation (Macaulay et al., 2014), mRNA expression profiles throughout meiosis
warrant investigation, as does investigation of oocyte-cumulus cell communications.
Hormone receptors
Exposure to 15 ng/mL BPA during oocyte maturation resulted in increased TRβ mRNA
expression in MII oocytes compared to the IVM group (Figure 14B). There were no significant
differences in ERβ mRNA expression as a result of BPA exposure in MII oocytes (Figure 14A).
Due to the presence of TRs in granulosa cells, and GV, MI, and MII oocytes, it has been suggested
that there may be a direct effect of THs on the maturing oocyte by interacting with thyroid
receptors in the surrounding cumulus and granulosa cells (Wakim et al., 1993; Zhang et al., 1997;
Zhang et al., 2007; Aghajanova et al., 2009; Costa et al., 2013). BPA has previously been found
to decrease TRβ levels in Xenopus embryos (Iwamuro et al., 2003), however to the best of our
knowledge, this is the first study analyzing the effects of BPA on TR levels following exposure
during oocyte maturation.
Our lab has previously shown that embryo treatment with TH improves embryo quality,
however when this treatment occurred at IVM no statistical differences were observed (Ashkar et
al., 2010b). Similar results have since been reported by Costa et al. (2013) where cleavage and
blastocyst rates were unaffected by IVM addition of THs, although increased blastocyst hatching
rates were observed (Costa et al., 2013). Furthermore, TR levels in the mature oocyte were
unaffected by TH supplementation (Ashkar et al., 2010b). In contrast we have observed alterations
in TRβ following BPA exposure which may suggest interaction with the receptor, or interference
with TRβ mRNA processing, resulting in altered expression in the MII oocyte. Therefore, BPA at
the 15 ng/mL dose appears to be acting as an endocrine disruptor in the current model.
114
E2 is known to play an important role in oocyte and embryo development, and levels can
improve developmental competence of in vitro matured oocytes (Kreiner et al., 1987; Zheng et al.,
2003; Modina et al., 2007; Aardema et al., 201), however elevated levels of E2 have also been
linked to poor developmental outcomes such as disruption to meiotic spindles (Beker-van
Woudenberg et al., 2004). Currently we observed no significant alterations in ER mRNA
expression in MII oocytes as a result of E2 or BPA exposure during oocyte maturation. Many nongenomic effects of BPA have been suggested such as interaction with membrane bound receptors
(Alonso-Magdalena et al., 2012), and epigenetic effects (Susiarjo et al., 2007). In the current
analysis BPA exposure at the doses tested does not influence ERβ mRNA expression in MII
oocytes or resulting blastocysts, and further investigation of BPA’s estrogenic effects, possibly
through non-genomic mechanisms, is required.
Although minor non-significant alterations in gene expression can be observed in the
blastocyst, it is apparent that exposure to BPA during oocyte maturation alone does not alter gene
expression of nuclear receptors (Figure 15). Thus new transcription arising from the embryo, at
least for the genes studies currently, is unaffected. Additional investigation of gene and protein
expression during early cleavage stages is important in further analysis of BPA’s influence on
hormone receptors in the early embryo. Though the current study was intended to observe effects
as a result of BPA exposure during oocyte maturation, it is established that gene expression in the
blastocyst is more likely to be altered as a result of environmental perturbations during embryonic
development. Therefore further analyses are required to assess exposure to BPA not only during
oocyte maturation, but also during embryonic development.
115
Stress and metabolism genes
In the current analysis, exposure to 15 ng/mL BPA during oocyte maturation resulted in
significantly higher p53 mRNA levels relative to the IVM and 30 ng/mL BPA groups in MII
oocytes (Figure 16). There were no significant changes in gene expression in the HSP70 and
GLUT1 mRNA expression levels. Due to the role that p53 plays in oxidative stress, DNA repair,
cell cycle regulation, and centrosome function, alterations in p53 expression could lead to
detrimental developmental effects in the embryo (Tchang & Méchali, 1999; Tritarelli et al., 2004;
Pascreau et al., 2009). Oxidative stress can result in loss of meiotic spindle integrity and errors in
chromosome segregation in oocytes, as well as increased apoptosis and decreased fertilization and
development rates (Tarin et al., 1996, Tatemoto et al., 2000; Takahashi et al., 2003; Tatone et al.,
2008; Vandaele et al., 2010). Additionally, p53 has been found to interact closely with AURKA
in Xenopus oocytes and mammalian cells (Pascreau et al., 2009), and disruption to p53 results in
similar phenotypes to that of AURKA such as centrosome amplification and aneuploidy (Pascreau
et al., 2009). AURKA and the cell cycle regulator, microtubule-associated TPX2 regulate the
phosphorylation of p53 (Pascreau et al., 2009). DAZL-directed translation during oocyte
maturation has also been found to interact with AURKA and p53 during oocyte maturation through
activation of various cell cycle regulators, such as TPX2 (Chen et al., 2011). Thus, the changes
observed in p53 in the current study may be linked to alterations observed in AURKA and DAZL
mRNA expression.
We found no significant alterations in mRNA levels of stress- and metabolism-related
genes at the blastocyst stage between treatment groups (Figure 17). The lack of alterations in
mRNA levels of these genes at the blastocyst stage indicates that embryos that survive to the
blastocyst stage following in vitro oocyte exposure to BPA are not exhibiting a stress response at
116
that time. Expression of these genes at early cleavage stages and as a result of embryonic exposure
is warranted to gain a fuller understanding of whether these genes are involved in BPA’s activities
during early development.
It is important to recognize that an ethanol effect was observed in many of the groups,
although this was only significant in the AURKA analysis. Thus although mRNA expression was
significantly higher in the 15 ng/mL BPA group than that of the IVM group, the ethanol in which
the BPA is dissolved is likely partly responsible for this increase. This increase in 15 ng/mL BPA
was observed in several genes, and a seemingly opposite effect in the higher BPA group relative
to the vehicle control, which although is not statistically significant, may indicate that BPA is
causing subtle alterations to a variety of genes. Subtle alterations such as these may be responsible
for larger effects such as changes in protein levels or epigenetic effects, and therefore these
parameters require further investigation.
The molecular events of oocyte maturation are still largely unclear, and how changes in
mRNA levels in the MII oocyte occur is not well understood. However it has been suggested that
environmental stress imposed on the maturing oocyte can alter mRNA storage and stability,
thereby affecting MII and embryonic gene expression (Gendelman & Roth, 2012). The alterations
in mRNA expression in MII oocytes observed in the current study may therefore indicate that BPA
can interfere with mechanisms responsible for gene processing, and may affect mRNA stability,
translation, and degradation, possibly through altering polyadenylation of specific genes.
Additional analyses are important to further investigate the molecular actions of BPA during
maturation.
117
GENERAL DISCUSSION
Female fertility can be affected by a variety of factors, including physical, environmental,
and lifestyle factors. The oocyte is at the center of a woman’s fertility and oocyte quality is critical
for normal fertilization, embryo development, and production of a healthy offspring. The oocyte
develops, grows, and matures within the ovarian follicle, and the follicular environment can be
altered by maternal health, lifestyle, and exposures. For instance, toxic exposures such as alcohol,
smoking, drugs, and various other chemicals may increase the risk of fertility issues (Mtango et
al., 2008, Varghese, 2010). Alterations in the composition of follicular fluid can not only lead to
poor oocyte quality, but can also result in a decreased embryo quality; thus, factors that can alter
normal follicular fluid concentrations, or enter the follicular fluid itself, are of great concern.
The main goal of this study was to identify if BPA exposure of bovine oocytes during in
vitro maturation leads to observable alterations in oocyte or embryo quality and developmental
potential. All exposures in the current thesis occurred only during the window of oocyte
maturation, thus all results observed are the consequence of alterations occurring during that period
in development. BPA has been shown in vitro and in vivo to negatively affect various aspects of
reproduction, and developmental exposure to the chemical has resulted in long-term, and even
trans-generational, effects (Cantonwine et al., 2010; Wei et al., 2011; Wolstenholme et al., 2012).
How BPA affects the mammalian, particularly the bovine, oocyte and early embryo is not as well
understood. Considering the detection of BPA in follicular fluid (Ikezuki et al., 2002), the effects
of BPA on the maturing oocyte are of great concern. Analysis of early parameters of developmental
competency, such as oocyte maturation and embryonic viability, may provide insight into how
developmental exposure prior to fertilization can disrupt early development and what implications
this may have for further development.
118
The environment in which the oocyte matures is critical to the developmental fate of the
oocyte, and can affect developmental capabilities of the subsequent embryo (Combelles &
Albertini, 2002). Oocytes are a long-lived cell population, however, nuclear maturation, the period
of time between the plasma LH surge and the oocyte’s rearrest at MII, is a critical window of
development for the establishment of oocyte competence (Lonergan et al., 2003). The composition
of follicular fluid is vital to and has the potential to affect oocyte maturation and quality as well as
embryo quality and viability. Appropriate hormone levels in follicular fluid and IVM media are
important in the acquisition of developmental competence (Kreiner et al., 1987; Zheng et al., 2003;
Modina et al., 2007; Aardema et al., 2013). In addition to abnormal hormonal levels,
environmental factors such as abnormal temperature, oxygen tension, in vitro culture conditions,
and glucose levels in follicular fluid, can disrupt oocyte maturation, resulting in decreased
embryonic developmental potential (Albertini & Carabatsos, 1998; Krisher & Bavister, 1999;
Spindler et al., 2000; Trounson et al., 2001; Hodges et al., 2002; reviewed by Krisher 2013). Since
oocyte competence is both critical to further developmental success and easily affected by
environmental factors, suboptimal environmental conditions of the oocyte during maturation may
have significant long-term effects on offspring arising from such oocytes.
Thus the current study investigated the effects of BPA exposure during in vitro oocyte
maturation on the quality and developmental potential of both oocytes and embryos, as well as
transcript levels of meiosis-, stress-, metabolism-, and hormone-related genes. The key findings of
this thesis are as follows. BPA treatment at 30 ng/mL resulted in an average uptake of 2.48 ng/mL
BPA per oocyte and led to decreased meiosis progression, increased MII spindle abnormalities,
decreased embryonic first cleavage, decreased blastocyst formation rate, increased apoptosis in
blastocysts and skewed sex ratio toward females. There were no significant gene expression
119
alterations in MII oocytes or blastocysts at this dosage level. The 15 ng/mL BPA treatment, which
resulted in decreased blastocyst development compared with the no-treatment control, led to
significantly higher p53 expression in MII oocytes than that resulting from 30 ng/mL BPA
treatment. Finally, the E2 treatment did not have any effects in comparison to the controls.
The experiments presented in Chapter 1 were conducted in order to assess the effects of
BPA exposure during oocyte maturation in vitro on meiosis progression and the MII spindle. The
stage of meiosis at 24 hours post IVM culture, and MII spindle morphology and organization were
assessed in order to determine BPA’s effects on oocyte quality and potential. Oocyte exposure to
30 ng/mL BPA resulted in an average uptake of 2.48 ng/mL per oocyte, and a decrease in oocyte
quality and developmental potential due to disruption of meiosis and induction of MII spindle
abnormalities. Due to the vulnerability of the oocyte to its microenvironment during maturation, a
suboptimal environment can interfere with developmental potential and hinder embryonic
development (van de Leemput et al., 1999; Sanfins et al., 2003; Ibáñez et al., 2005). Abnormalities
occurring during oocyte maturation can decrease oocyte quality and developmental potential and
can therefore compromise future embryonic development (Hyttel et al., 1989; Sirard et al., 2006).
The meiotic spindle, a key component of nuclear maturation, is a microtubular structure
that acts primarily to assist in chromatid segregation and the associated second polar body
extrusion occurring at the conclusion of meiosis (Coticchio et al., 2010). The meiotic spindle is
exceptionally vulnerable to environmental factors such as oxidative stress (Hu et al., 2001;
Eichenlaub-Ritter et al., 2002), which can induce meiotic abnormalities and chromosome
instability, and lead to impaired embryo development and increased apoptosis (Liu et al., 2003) as
was observed in the current study. The MII spindle can be adversely affected by factors such as
changes in pH (Swain, 2010), high E2 levels (Beker et al., 2002; Beker-van Woudenberg et al.,
120
2004), and exposure to EDCs (Cecconi et al., 2007). Flattening of the spindle poles has been
associated with increased chromosome misalignment (Coticchio et al., 2013; Ferris et al., 2015;
Chapter 1). Chromosome misalignment may lead to improper chromosome segregation, which is
relatively common in human oocytes (Hassold & Hunt, 2001). Though aneuploidy is common in
humans, the proportion of aneuploidy cells within an embryo directly correlates with
developmental potential (Baltaci et al., 2006; Mtango et al., 2008), and these errors have been
linked to pregnancy failure, embryonic abnormalities, and chromosomal disorders (Hassold &
Hunt, 2001; Mtango et al., 2008; Li & Albertini, 2013).
Meiosis progression, spindle morphology, and chromosome alignment were all negatively
altered by exposure to 30 ng/mL BPA during maturation. This has been observed previously in
mouse and human models (Hunt et al., 2003; Can et al., 2005; Susiarjo et al., 2007; EichenlaubRitter et al., 2008), and BPA has been linked to embryonic aneuploidy in some (Hunt et al., 2003;
Susiarjo et al., 2007), but not other experimental studies (Eichenlaub-Ritter et al., 2008). The
disruption of meiosis progression and spindle organization as a result of BPA exposure during
oocyte maturation indicates both unsuitable environmental conditions for the oocyte during this
time, as well as an increased possibility of further developmental deficiencies. The current studies
show that BPA at a sufficient concentration can be taken up by the oocyte and lead to disruption
of meiosis and interference with normal MII spindle organization and chromosome alignment.
The mechanisms of spindle disruption are not clear, but oocyte-somatic cell
communication is critical to spindle integrity. Disruptions to this communication may interfere
with normal oocyte maturation (Albertini et al., 2001; Hodges et al., 2002). Furthermore, meiosis
may be affected by BPA through disruption of the microtubules directly, and by interaction with
proteins associated with the microtubules (Pfeiffer et al., 1997; Can et al., 2005). Meiosis
121
disruption may be due to selective interference with centrosome and microtubule organization
(Can et al., 2005). Unlike other estrogenic agents that can disrupt meiosis through
depolymerisation of microtubules, the actions of BPA appear to be a result of disorganization and
fragmentation of centrosomes, resulting in spindle disorganization (Can et al., 2005). Motor
proteins and regulatory mechanisms of transportation associated with the microtubules may be
target sites for BPA during oocyte maturation. For instance, BPA may lead to errors in cell cycle
progression, microtubule organization and centrosome assembly and function by interfering with
the interaction between protein kinase C (PKC) and pericentrin, interfering with centrosomal
proteins, disrupting protein transport, or by the disorganization of the microtubule organizing
centers (MTOCs) (Takahashi et al., 2000; Can et al., 2005).
Because there appears to be a correlation between meiosis progression and spindle
organization and later embryonic development, the present study assessed developmental
parameters such as development rate, total cell number, apoptosis, sex ratio, and gene expression
to evaluate oocyte and embryo quality and developmental potential. Oocyte and embryo quality
are directly related, and poor quality oocytes exhibit decreased fertilization success, and embryo
quality (Metwally et al., 2007). Although the embryonic environment has a great impact to the
developmental fate of the embryo, oocyte quality can dictate the development and survival of an
embryo, pregnancy establishment and maintenance, and fetal development (reviewed by Varghese
et al., 2010). Taking into account that follicular fluid can alter the quality and developmental
potential of oocytes, the results presented in Chapter 2 therefore indicate that oocyte exposure to
BPA at certain concentrations can impact development of the preimplantation embryo, likely by
decreasing quality of the mature oocyte. Cleavage and blastocyst rates were decreased, apoptosis
122
rate, as well as the proportion of cells with non-apoptotic DNA fragmentation, was increased, and
there was a higher proportion of female embryos at blastocyst.
Oxidative stress, heat stress, abnormal hormone levels, in vitro culture conditions,
abnormal oocyte-somatic cell signalling and exposure to toxic or xenobiotic chemicals can lead to
decreased developmental competency of the embryo (Albertini & Carabatsos, 1998; Betts & King,
2001; Trounson et al., 2001; Hodges et al., 2002; Hamdoun & Epel, 2007). The effects of stress
and/or abnormal hormone levels during preimplantation development have been linked to
decreased development, increased apoptosis (Paula-Lopes & Hansen, 2002b), and skewed sex
ratio (Krackow, 1997), but the evidence of embryonic effects as a result of oocyte impairments is
not as well studied. Altered meiotic success and spindle abnormalities have been linked to a variety
of developmental effects, however it is unclear if it is these meiotic changes themselves that
influence further development, for instance by way of improper chromosome segregation leading
to aneuploidies in embryos, or through other pathways that affect both maturation and further
embryonic development such as poor oocyte-cumulus cell communication.
As mentioned previously, several studies indicating oocyte effects such as decreased
meiosis success and spindle abnormalities also resulted in poor embryonic outcomes (van de
Leemput et al., 1999; Ertzeid & Storeng, 2001; Van der Auwera & D’Hooghe, 2001; Sirard et al.,
2006). Meiotic abnormalities have been linked to later developmental effects such as impaired
fertilization, embryo development, and cell cycle progression as well as an increased incidence of
apoptosis in blastomeres, embryonic abnormalities, and pregnancy loss (Volarcik et al., 1998;
Sanfins et al., 2003; Cecconi et al., 2007; Bromfield et al., 2009). Furthermore, early cleaving
embryos, which are considered to have greater developmental potential than later cleaving
embryos, have been shown to have a greater spindle size and higher blastocyst formation rate
123
whereas later cleaving embryos exhibited a smaller, compressed spindle and a lower blastocyst
rate (Tomari et al., 2011). Meiotic spindle parameters are strong indicators of human embryo
developmental potential, thus the decreased development rates currently observed following
oocyte exposure to BPA could be directly related to the meiotic disturbances that were presented
in Chapter 1. The mechanism by which this may occur is not clear; however, alterations in genes
and protein expression, oocyte metabolism, and communication between the oocyte and
surrounding cells may play a key roles in the observed effects (Van Blerkom et al., 1995; Tomari
et al., 2011).
Embryos have been shown to respond to stress by undergoing apoptosis (Byrne et al, 1999).
Apoptotic cell death in early embryos has been found to occur in a variety of mammalian species
under certain stressful conditions once the genome of embryo has been activated. The degree of
apoptotic response plays a role in the embryo’s ability to continue development. A limited
apoptotic response allows the embryo to continue development by removing damaged cells
without detrimentally affecting neighbouring cells. More extensive apoptosis may result in the
removal of a relatively large proportion of cells thereby lowering the embryo’s viability or chance
of survival (Betts & King, 2001; Paula-Lopes & Hansen, 2002b). Depending on the conditions and
severity of the stress encountered, apoptosis may serve as a marker of an embryo at risk and/or as
a mechanism to help the embryo survive suboptimal conditions. As was presented in Chapter 2,
preimplantation embryos exhibited increased apoptosis under the same treatment conditions that
resulted in increased meiotic abnormalities.
An altered sex ratio of blastocyst may also indicate stress experienced by the embryo
(Krackow, 1997; Pratt & Lisk, 1989; Jiménez et al., 2003). The blastocyst sex-ratio was skewed
towards females in both BPA groups, however this was only significantly different than the
124
expected 1:1 ratio in the 30 ng/mL BPA group (Chapter 2). This may indicate that the embryos
experienced heightened stress, resulting in the disproportionate arrest of male embryos.
Differences in how female and male embryos respond to environmental conditions are speculated
to be a result of metabolic, genetic, and epigenetic differences. During the preimplantation period,
male and female embryos differ only in the content of their sex chromosomes, and differences
exhibited by embryos at this time may be a result of transcriptional dimorphism (Bermejo-Alvarez
et al., 2011). Furthermore, epigenetic differences resulting from the presence of one versus two Xchromosomes may be a driving force in sex differences and how embryos respond to
environmental conditions (reviewed by Gutiérrez-Adán et al., 2006).
Expression of genes encoded by the sex chromosomes, which may also affect autosomal
gene expression, differs between male and female preimplantation embryos (reviewed by
Bermejo-Alvarez et al., 2011). Y-linked genes are only expressed in male embryos, and X-linked
genes are expressed doubly in females. X-chromosome inactivation (XCI) compensates for this
disparity ensuring X-linked genes (in most cases) are transcribed equally in male and female adult
tissues (reviewed by Bermejo-Alvarez et al 2011). However, if during early embryo development
XCI is incomplete, or genes are reactivated, the female embryos exhibit higher expression of many
X-linked genes (Kobayashi et al., 2006; Bermejo-Alvarez et al. 2010a). This phenomenon has been
found in mouse (Kobayashi et al., 2006), bovine (Gutiérrez-Adán et al., 2000), and human embryos
(Taylor et al., 2001).
These transcriptional alterations are responsible for variations between male and female
embryos such as metabolic differences, and dimorphic susceptibilities to suboptimal in vivo and
in vitro conditions (reviewed by Gutiérrez-Adán et al., 2006). Resulting alterations in molecular
pathways, such as the pentose-phosphate pathway (PPP) that regulates glucose metabolism, may
125
result in varying susceptibilities of male and female embryos. For instance, male embryos have
been reported to metabolize glucose at a higher rate than females (Tiffin et al., 1991) and male
embryos appear to be more developmentally competent than females under hyperglycemic
conditions in vitro, however the opposite has also been found (Jimenez et al., 2003). Conversely,
female embryos exhibit increased expression of X-linked genes such as those related to energetic
metabolism, the regulation of oxygen radicals, and apoptosis inhibition (Gutiérrez-Adán et al.,
2000; Jimenez et al., 2003). Since various stressors can lead to embryonic overproduction of
reactive oxygen species (ROS), female embryos may be better equipped to survive such a stress
due to enhanced ability to buffer the amount of cellular ROS (Perez-Crespo et al., 2005).
Furthermore, male embryos have been shown to be more sensitive to oxidative damage induced
by heat stress, which may be due to the increased expression of X-linked genes in females that
result in an increased ability to buffer environmental stress (Perez-Crespo et al., 2005).
Therefore, whether the embryo possesses one or two X chromosomes may underlie the
differences in the early embryo (Gutiérrez-Adán et al., 2006). This may explain observations that
male embryos may be more vulnerable to stressors than female embryos under certain
environmental conditions. Since male and female embryos evidently respond differently to stress,
a skewed sex ratio may be indicative of environmental stress imposed on the embryo. Therefore,
the skewed sex ratio observed in the present studies may indicate a stress response activated in the
embryos resulting from oocytes exposed to 30 ng/mL BPA during maturation. Taken together, the
results presented in Chapter 2 suggest that embryo quality and developmental potential were
decreased. The decrease in quality as evidenced by increased apoptosis and a skew in sex ratio
indicates that adverse environmental conditions may have elicited a stress response in the embryo.
126
Chapters 1 and 2 presented evidence that BPA exposure during oocyte maturation
decreased oocyte and embryo quality and developmental potential. BPA has previously been
shown to affect transcript levels of a number of genes. Hormone receptors (Rubin, 2011), cell cycle
regulators (Peretz et al., 2012), apoptotic genes (Peretz et al., 2012), genes related to the stress
response (Tabuchi et al., 2002), and those related to the onset of meiosis, chromatin modification,
remodeling, and chromosome condensation (Lawson et al., 2011), among others have been altered
as a result of BPA exposure. To determine if BPA was interfering with spindle-related gene
processing, inducing embryonic stress, or altering hormone pathways, genes involved in meiosis
and spindle assembly (CDC2, AURKA, DAZL, KIF5b), stress and metabolism (HSP70, p53, and
GLUT1), and genes of nuclear hormone receptors (ERβ, and TRβ), were assessed at the oocyte
and/or blastocyst stage. Alterations in mRNA levels in MII oocytes may indicate poor
developmental competency and could result in poor developmental outcomes (Paczowski et al.,
2011; Yuan et al., 2011). Blastocysts are a key developmental milestone in early embryonic
development, and alterations in mRNA abundance at this stage may indicate an altered quality, or
developmental competency of the blastocyst (reviewed by Duranthon et al., 2008). Thus, Chapter
3 consisted of mRNA quantification of genes involved in meiosis, embryonic stress, metabolism,
and hormone-signalling in oocytes and blastocysts.
Oocyte maturation consists of a variety of tightly regulated sequential events mediated by
transient production and phosphorylation of regulatory proteins (reviewed by Pelech et al., 2008;
Brunet & Verlhac, 2011). Kinases and phosphatases act to initiate and maintain proper sequential
activities throughout oocyte maturation (Yamasita et al., 2000). Meiotic progression and cell
division are mediated by the selective and specific regulation of cellular components by translation
and protein synthesis, cytoplasmic polyadenylation, phosphorylation, and protein degradation
127
(Richter, 1991, 2007; reviewed by Uzbekova et al., 2008; Chen et al., 2011). Normal processing
of genes involved in cell cycle progression and spindle-assembly is critical to the ability of the
oocyte to acquire developmental competence (Tanaka et al., 1998; Takamiya et al., 2004;
Uzbekova et al., 2008; Lane et al., 2010; Chen et al., 2011). Considering the disruption to meiosis
as well as the spindle abnormalities observed in Chapter 1, several genes important to meiotic cell
cycle progression and spindle assembly and integrity were analyzed in MII oocytes following
maturation under various treatment conditions.
Treatment with 15 ng/mL BPA resulted in increased expression of CDC2, AURKA, DAZL
in MII oocytes. There appeared to be a slight vehicle effect, with the vehicle group (0.1% ethanol)
resulting in significantly increased expression of AURKA mRNA and non-significant increases in
several other genes analyzed. Similar gene expression patterns were observed in MII oocytes of
CDC2, AURKA, and KIF5b between treatment groups, characterized by non-significant decreases
in gene expression in the E2 and the 30 ng/mL BPA groups, and no change or increased expression
in the 15 ng/mL BPA group relative to the vehicle control (0.1% ethanol). Thus although some
non-significant decreases were observed in the E2 and 30 ng/mL BPA groups compared to the
vehicle control, considering the increase in gene expression and lack of developmental effects
observed as a result of ethanol exposure, the functional significance of these observations cannot
be ascertained.
BPA has been shown to interact with the ERs and TRs (Matthews et al., 2001; Kang et al.,
2002; Moriyama et al., 2002; Iwamuro et al., 2003; Takao et al., 2003; Levy et al., 2004;
Lahnsteiner et al., 2005; Zoeller et al., 2005; Vandenberg et al., 2009; Meeker & Ferguson, 2011).
BPA has caused alterations in mRNA levels of these genes in reproductive and non-reproductive
tissues (Tabuchi et al., 2002; Ramos et al., 2003; Markey et al., 2005; Berger et al., 2010;
128
Aghajanova & Giudice, 2011). In Chapter 3 the lower dose of BPA (15 ng/mL; 1.69 ng/mL
BPA/oocyte), resulted in an increased TRβ mRNA expression in MII oocytes compared to the
IVM group, and no effects were observed in ERβ mRNA expression in MII oocytes. There were
no significant alterations in either ERβ or TRβ in blastocysts. Alterations in TRβ following BPA
exposure may suggest interaction with the receptor, or interference with TRβ mRNA processing,
resulting in altered expression in the MII oocyte. Therefore, BPA at the 15 ng/mL dose appears to
be acting as an endocrine disruptor in the current model, however these results must be interpreted
with caution due to the vehicle effect observed. Further analysis of BPA’s non-genomic effects, as
well as analysis of exposure during embryonic development, may further help to elucidate
hormone-disrupting properties of BPA in the current model.
Due to the embryonic effects observed in Chapter 2, it was decided to evaluate stressrelated genes to see if transcripts of stress-related genes were altered in MII oocytes, and if a stressresponse was induced in the blastocyst. Exposure to 15 ng/mL BPA during oocyte maturation
resulted in significantly higher p53 mRNA levels relative to the IVM and 30 ng/mL BPA groups
in MII oocytes. There were no differences in HSP70 and GLUT1 mRNA expression levels in MII
oocytes. The significant difference observed between the two BPA groups is interesting and
supports the idea that the effects induced by BPA differ according to the exposure doses
administered. Given that p53 and AURKA are known to interact during oocyte maturation
(Pascreau et al., 2009), the results presented indicate a disruption to this molecular pathway. No
significant alterations were found in any of the groups for the three genes (HSP70, p53, and
GLUT1) in the blastocyst stage, suggesting that new transcription following the MET may not be
affected by BPA exposure at the oocyte stage. However earlier stages both before and during the
MET are necessary to determine whether BPA can alter mRNA expression of these genes.
129
Alterations observed in p53 mRNA in MII oocytes by BPA treatment may indicate that
BPA is inducing oxidative stress. Oxidative stress is thought to be a common mechanism by which
endocrine disruptors exert their effects (Wang et al., 2012), and is also a well characterized
example of how oocyte stress can affect embryonic development (reviewed by Guérin et al., 2001).
Stress experienced during oocyte maturation can lead to the production of ROS in embryo
development and lead to alterations in embryonic gene expression (Maître et al., 1993; Schultz,
1993; Guérin et al., 2001). The mRNA transcripts of various antioxidant enzymes have been found
in human, bovine, and murine GV and MII oocytes (El Mouatassim et al., 1999), indicating their
importance for further development (Harvey et al., 1995; reviewed by Guérin et al., 2001). Due to
the correlation between mRNA, proteins and enzyme activity levels of antioxidant enzymes, it has
been suggested that these enzymes are primarily regulated at the pre-translational level (Guérin et
al., 2001). Therefore, since maternal transcripts decrease until the maternal to embryonic transition
(MET), variations in the synthesis or accumulation of mRNAs of antioxidant enzymes during
oocyte maturation may impact embryonic development (Guérin et al., 2001). Thus, the
developmental effects observed in Chapter 2, taken together with the current findings may indicate
that BPA is affecting oocyte quality and developmental potential resulting in abnormal early
embryonic development. Further analysis of early cleavage stages is required to expand on this
observation.
All of the effects discussed in this thesis must be brought back to the oocyte. Since exposure
only occurred over a short period of time during in vitro oocyte maturation, key events occurring
during that window set the stage for all of the results observed in the current study. How BPA
exerts its effects at the oocyte may provide key insight into the mechanism by which the embryonic
effects are elicited. Clearly BPA affects oocyte quality, which can lead to developmental effects
130
in the preimplantation embryo. But exactly how these oocyte and embryonic effects are elicited is
difficult to determine. The developmental effects observed in the oocyte and embryo following
BPA exposure during oocyte maturation may be due to a number of mechanisms. Interfering with
microtubule assembly, interruption of proper metabolism, epigenetic alterations, improper
degradation of maternal mRNAs and proteins, and interference with communication between
cumulus cells and the oocyte, can all lead to interruptions to normal oocyte and embryonic
development (Pfeiffer et al., 1997; Hassold & Hunt, 2001; Can et al., 2005; Wetherill et al., 2007;
Lenie et al., 2008).
The communication between the oocyte and cumulus cells may be a key factor involved in
BPA’s effects. This bidirectional signaling is critical to oocyte development, maturation, and the
acquisition of developmental competence (reviewed by Hassold & Hunt, 2008). The oocyte drives
development by secreting factors that influence the growth, proliferation, and differentiation of its
surrounding cells, to which it depends on for growth and competence (Eppig, 2001; Sugiura et al.,
2005; Diaz et al., 2006, 2007; Mtango et al., 2008). Furthermore, the cumulus cells regulate
meiosis (Chesnel et al., 1994), and modulate oocyte genomic transcriptional activity (De La Fuente
& Eppig, 2001). Disruptions to these processes can result in reduced ovulation, fertilization and
developmental potential (Yan et al., 2001; Di Pasquale et al., 2004). BPA exposure may interfere
with these communications, altering the quality and developmental potential of both oocytes and
resulting embryos.
Oocyte maturation is an intricate event and alterations in this process can lead to poor
developmental outcomes down the line. Factors such as the xenoestrogen BPA can contaminate
follicular fluid and disrupt oocyte maturation and are thus of great concern. Although many
reproductive effects of BPA have been reported, the mechanism of action of BPA on oocyte
131
maturation is still being elucidated. Although E2 has been shown to impair meiotic progression
and spindle morphology in some cases during in vitro maturation (Beker-van Woudenberg et al.,
2004), in the current study E2 did not show any detrimental effects of oocyte maturation and
embryo development. BPA may not be acting as an estrogen mimic in the current analysis, but it
appears that BPA is acting, at least in part, as an endocrine disruptor, possibly by interfering with
endocrine pathways in ways such as activation of hormone receptors or alterations in hormone
metabolism. Disruptions to normal oocyte maturation may influence short- and long-term viability.
The mechanisms by which environmental factors influence oocyte maturation are important in
order to determine the risk of exposure and how chronic exposure could impact long-term fertility.
132
SUMMARY AND CONCLUSIONS
Disruption to the environment in which an oocyte matures can result in short- and longterm developmental effects. Exposure to 30 ng/mL BPA during oocyte maturation resulted in an
average uptake levels of 2.48 ng/mL per oocyte, and an associated disruption to oocyte maturation,
compromised oocyte and embryo quality and reduced developmental potential.
Exposure to a BPA concentration of 30 ng/mL during oocyte maturation decreased meiosis
success and increased the incidence of MII spindle abnormalities including an abnormally shaped
spindle (compressed with loss of pole focus) as well as chromosome misalignment.
Oocyte exposure to 30 ng/mL BPA decreased embryo cleavage and development rates and
increased the proportion of apoptotic blastomeres and nuclei containing non-apoptotic DNA
damage in the blastocyst, and caused a sex skew towards femaleness. The 15 ng/mL BPA group
had decreased blastocyst formation rate in comparison to the no-treatment control. Total cell
number did not differ between any of the groups.
Treatment with 15 ng/mL BPA resulted in increased expression of CDC2, AURKA,
DAZL, TRβ and p53 in MII oocytes relative to that of the IVM group. This increased p53
expression resulting from the 15 ng/mL BPA group was also significantly greater than the
expression levels resulting from 30 ng/mL BPA. There appeared to be a slight vehicle effect, with
the vehicle group (0.1% ethanol) resulting in significantly increased expression of AURKA mRNA
and non-significant increases in several other genes analyzed. There were no significant
differences observed in blastocyst mRNA expression levels of the genes analyzed.
The results obtained support the working hypothesis of the study that exposure to BPA
during oocyte maturation can, in a dose-dependent way, decrease oocyte and embryo quality and
133
developmental potential. Exposure of bovine oocytes in vitro to 30 ng/mL BPA during maturation
induced meiotic perturbations as well as poor embryonic outcomes including decreased
development rates, increased apoptosis and DNA fragmentation, and a skewed sex ratio, however
did not result in significant changes in mRNA expression relative to controls. Although gene
expression was not affected at this dose, it is clear that exposure to BPA only during in vitro
maturation can induce effects that are observable in the blastocyst.
134
FUTURE DIRECTIONS
Since the current study was the first, to our knowledge, to assess BPA exposure during
oocyte maturation in a bovine model, much of the thesis was dedicated to whether or not there was
an observable effect in the oocyte as well as in the subsequent embryo. Further analyses of gene
expression as a result of embryonic exposure to BPA would clarify if the blastocyst is more
vulnerable to embryonic exposures rather than those at the oocyte stage.
Oocyte maturation can be further assessed by analyzing temporal gene transcript and
protein levels for genes essential to developmental competence, those involved in oocyte-cumulus
cell communication, as well as genes that have been altered as a result of oocyte stress such as
those of antioxidant enzymes. Analysis of mRNA stability and degradation during oocyte
maturation will help to determine if BPA is altering mRNA expression by disrupting transcript
degradation. Because of the importance of communication between cumulus cells and the oocyte
during maturation, evaluation of whether BPA interrupts or alters signalling during maturation
may provide key understanding of the disruptive capabilities of BPA at this time. Additionally,
looking at transcriptional and translational alterations in the cumulus cells during maturation is
likely critical in the evaluation of communication within the COC.
Embryo developmental parameters measured can be expanded by observing earlier
developmental effects such as fertilization success, the stage and sex of arrested embryos,
apoptosis levels during the MET, embryonic aneuploidy, and further analysis of stress and
hormonal pathways. Co-supplementation of IVM media with E2 or THs as well as antagonists
may provide information on whether BPA is acting through E2 or TH pathways. Furthermore,
incubation with antioxidants or HSP antibodies may provide further information as to whether
maturational exposure to BPA is resulting in stress-related effects. Alterations in both oocyte and
135
embryonic metabolism may also be important in gaining a greater understanding of the effects of
BPA on early developmental stages.
Further analysis of gene and protein expression at the blastocyst as well as earlier in
development will continue to provide insight into the developmental effects of BPA. Analysis at
all stages of development would also help to provide information on whether BPA affects maternal
transcripts, embryonic, or both and when the greatest disruption is. Protein expression during
embryo development would also provide key information on how these alterations lead to
functional differences in the early embryo. Furthermore, epigenetic analysis, such as methylation
levels of genes, will be key to understanding how early developmental effects can be linked to the
later developmental effects observed in offspring, adults, and the subsequent generation. Further
analysis of the delicate interactions during oocyte maturation is required to further elucidate the
mechanism by which BPA is affecting meiosis, and how this may be linked to the embryonic
effects observed.
136
REFERENCES
Aardema, H., Roelen, B. A. J., van Tol, H. T. A, Oei, C. H. Y., Gadella, B. M., & Vos, P. L. A
M. (2013). Follicular 17β-estradiol and progesterone concentrations and degree of cumulus
cell expansion as predictors of in vivo-matured oocyte developmental competence in
superstimulated heifers. Theriogenology, 80(6), 576–583.
doi:10.1016/j.theriogenology.2013.05.025
Aghajanova, L., & Giudice, L. C. (2011). Effect of bisphenol A on human endometrial stromal
fibroblasts in vitro. Reproductive Biomedicine Online, 22(3), 249–56.
doi:10.1016/j.rbmo.2010.12.007
Akingbemi, B. T., Sottas, C. M., Koulova, A. I., Klinefelter, G. R., & Hardy, M. P. (2004).
Inhibition of testicular steroidogenesis by the xenoestrogen bisphenol A is associated with
reduced pituitary luteinizing hormone secretion and decreased steroidogenic enzyme gene
expression in rat Leydig cells. Endocrinology, 145(2), 592–603. doi:10.1210/en.2003-1174
Albertini, D. F., Combelles, C. M. H., Benecchi, E., & Carabatsos, M. J. (2001). Cellular basis
for paracrine regulation of ovarian follicle development. Reproduction, 121, 647–653.
doi:10.1530/reprod/121.5.647
Albertini, D. F., Sanfins, A., & Combelles, C. M. (2003). Origins and manifestations of oocyte
maturation competencies. Reproductive BioMedicine Online, 6(4), 410–415.
doi:10.1016/S1472-6483(10)62159-1
Al-Katanani, Y. M., Paula-Lopes, F. F., & Hansen, P. J. (2002). Effect of season and exposure to
heat stress on oocyte competence in Holstein cows. Journal of Dairy Science, 85, 390–396.
doi:10.3168/jds.S0022-0302(02)74086-1
Alonso-Magdalena, P., Vieira, E., Soriano, S., Menes, L., Burks, D., Quesada, I., & Nadal, A.
(2010). Bisphenol a exposure during pregnancy disrupts glucose homeostasis in mothers
and adult male offspring. Environmental Health Perspectives, 118(9), 1243–1250.
doi:10.1289/ehp.1001993
Aluru, N., Leatherland, J. F., & Vijayan, M. M. (2010). Bisphenol A in oocytes leads to growth
suppression and altered stress performance in juvenile rainbow trout. PloS One, 5(5),
e10741. doi:10.1371/journal.pone.0010741
Amaral, J. D., Xavier, J. M., Steer, C. J., & Rodrigues, C. M. (2010), The role of p53 in
apoptosis. Discovery Medicine, 9(45), 145-152.
Anderson, R. L. (1998). Stress proteins and apoptosis in prenatal development, cancer and
medicine. Cell Stress & Chaperones, 3(4), 209-212.
Andrésson, T., & Ruderman, J. V. (1998). The kinase Eg2 is a component of the Xenopus oocyte
progesterone-activated signaling pathway. EMBO Journal, 17(19), 5627–5637.
doi:10.1093/emboj/17.19.5627
137
Aoki, T. & Takada, T., (2012). Bisphenol A modulates germ cell differentiation and retinoic acid
signaling in mouse ES cells. Reproductive Toxicology, 34(3), 463–470.
doi:10.1016/j.reprotox.2012.06.001.
Arck, P. (2004). Stress and embryo implantation. Journal de gynécologie, obstétrique et biologie
de la reproduction (Paris), 33, S40–S42
Ashkar F., Semple, E., Schmidt, C. H., St John, E., Bartlewski, P. M., & King, W. a. (2010b).
Thyroid hormone supplementation improves bovine embryo development in vitro. Human
Reproduction (Oxford, England), 25(2), 334–44. doi:10.1093/humrep/dep394
Ashkar, F. (2013). Developmental and genomic aspects of thyroid hormones during early
embryo development in cattle. PhD thesis. The University of Guelph.
Ashkar, F. a, Bartlewski, P. M., Singh, J., Malhi, P. S., Yates, K. M., Singh, T., & King, W. A.
(2010a). Thyroid hormone concentrations in systemic circulation and ovarian follicular
fluid of cows. Experimental Biology and Medicine (Maywood, N.J.), 235(2), 215–21.
doi:10.1258/ebm.2009.009185
Assou, S., Anahory, T., Pantesco, V., Le Carrour, T., Pellestor, F., Klein, B. Reyftmann, L.,
Dechaud, H., De Vos, J., & Hamamah, S. (2006). The human cumulus-oocyte complex
gene-expression profile. Human Reproduction, 21(7), 1705–1719.
doi:10.1093/humrep/del065
Avissar-Whiting, M., Veiga, K. R., Uhl, K. M., Maccani, M. A, Gagne, L. A, Moen, E. L., &
Marsit, C. J. (2010). Bisphenol A exposure leads to specific microRNA alterations in
placental cells. Reproductive Toxicology (Elmsford, N.Y.), 29(4), 401–6.
doi:10.1016/j.reprotox.2010.04.004
Bachvarova, R., De Leon, V., Johnson, a, Kaplan, G., & Paynton, B. V. (1985). Changes in total
RNA, polyadenylated RNA, and actin mRNA during meiotic maturation of mouse oocytes.
Developmental Biology, 108(2), 325–331. doi:10.1016/0012-1606(85)90036-3
Baker, M. E., & Chandsawangbhuwana, C. (2012). 3D Models of MBP, a Biologically Active
Metabolite of Bisphenol A, in Human Estrogen Receptor α and Estrogen Receptor β. PLoS
ONE, 7(10), 1–15. doi:10.1371/journal.pone.0046078
Balasubramanian, S., Son, W. J., Kumar, B. M., Ock, S. a, Yoo, J. G., Im, G. S., … Rho, G. J.
(2007). Expression pattern of oxygen and stress-responsive gene transcripts at various
developmental stages of in vitro and in vivo preimplantation bovine embryos.
Theriogenology, 68(2), 265–75. doi:10.1016/j.theriogenology.2007.05.044
Baldwin, S. A. (1993). Mammalian passive glucose transporters: Members of an ubiquitous
family of active and passive transport proteins. Biochimica et Biophysica Acta - Reviews
on Biomembranes, 1154, 17–49. doi:10.1016/0304-4157(93)90015-G
138
Baltaci, V., Satiroglu, H., Kabukçu, C., Unsal, E., Aydinuraz, B., Uner, O., … Aktan, a. (2006).
Relationship between embryo quality and aneuploidies. Reproductive Biomedicine Online,
12(1), 77–82. doi:10.1016/S1472-6483(11)60369-6
Batt, P. A., Gardner, D. K., & Cameron, A. W. (1991). Oxygen concentration and protein source
affect the development of preimplantation goat embryos in vitro. Reproduction, Fertility
and Development, 3(5), 601-607.
Battaglia, D. E., Goodwin, P., & Klein, N. A. (1996). Influence of maternal age on meiotic
spindle assembly in oocytes from naturally cycling women, 11(10), 2217–2222.
Baumann, C., Viveiros, M. M., & De La Fuente, R. (2010). Loss of maternal ATRX results in
centromere instability and aneuploidy in the mammalian oocyte and pre-implantation
embryo. PLoS Genetics, 6(9), e1001137. doi:10.1371/journal.pgen.1001137
Beker van Woudenberg, A. R., van Tol, H. T. A, Roelen, B. A. J., Colenbrander, B., & Bevers,
M. M. (2004). Estradiol and its membrane-impermeable conjugate (estradiol-bovine serum
albumin) during in vitro maturation of bovine oocytes: effects on nuclear and cytoplasmic
maturation, cytoskeleton, and embryo quality. Biology of Reproduction, 70(5), 1465–74.
doi:10.1095/biolreprod.103.025684
Beker van Woudenberg, A., Gröllers-Mulderij, M., Snel, C., Jeurissen, N., Stierum, R., &
Wolterbeek, A. (2012). The bovine oocyte in vitro maturation model: a potential tool for
reproductive toxicology screening. Reproductive Toxicology (Elmsford, N.Y.), 34(2),
251–60. doi:10.1016/j.reprotox.2012.05.098
Beker, a R. C. L., Colenbrander, B., & Bevers, M. M. (2002). Effect of 17beta-estradiol on the in
vitro maturation of bovine oocytes. Theriogenology, 58(9), 1663–73. Retrieved from
http://www.ncbi.nlm.nih.gov/pubmed/12472137
Berger, R. G., Foster, W. G., & deCatanzaro, D. (2010). Bisphenol-A exposure during the period
of blastocyst implantation alters uterine morphology and perturbs measures of estrogen and
progesterone receptor expression in mice. Reproductive Toxicology (Elmsford, N.Y.),
30(3), 393–400. doi:10.1016/j.reprotox.2010.06.006
Bermejo-Alvarez, P., Rizos, D., Lonergan, P., & Gutierrez-Adan, A. (2011). Transcriptional
sexual dimorphism during preimplantation embryo development and its consequences for
developmental competence and adult health and disease. Reproduction (Cambridge,
England), 141(5), 563–70. doi:10.1530/REP-10-0482
Bermejo-Alvarez, P., Rizos, D., Rath, D., Lonergan, P., & Gutierrez-Adan, A. (2010). Sex
determines the expression level of one third of the actively expressed genes in bovine
blastocysts. Proceedings of the National Academy of Sciences of the United States of
America, 107, 3394–3399. doi:10.1073/pnas.0913843107
Bertolini, M., Beam, S. W., Shim, H., Bertolini, L. R., Moyer, A. L., Famula, T. R., & Anderson,
G. B. (2002). Growth, development, and gene expression by in vivo- and in vitro-produced
139
day 7 and 16 bovine embryos. Molecular Reproduction and Development, 63(May), 318–
328. doi:10.1002/mrd.90015
Betteridge, K. J. (2001). Comparative aspects of conceptus growth: A historical perspective.
Reproduction, 122(September 2000), 11–19. doi:10.1530/reprod/122.1.11
Betts, D. H., & King, W. A. (2001). Genetic regulation of embryo death and senescence.
Theriogenology, 55(00), 171–191. doi:10.1016/S0093-691X(00)00453-2
Beydoun, H. a., Khanal, S., Zonderman, A. B., & Beydoun, M. A. (2014). Sex differences in the
association of urinary bisphenol-A concentration with selected indices of glucose
homeostasis among U.S. adults. Annals of Epidemiology, 24, 90–97.
doi:10.1016/j.annepidem.2013.07.014
Bloom, M. S., Kim, D., Vom Saal, F. S., Taylor, J. A, Cheng, G., Lamb, J. D., & Fujimoto, V. Y.
(2011a). Bisphenol A exposure reduces the estradiol response to gonadotropin stimulation
during in vitro fertilization. Fertility and Sterility, 96(3), 672–677.e2.
doi:10.1016/j.fertnstert.2011.06.063
Bloom, M. S., Vom Saal, F. S., Kim, D., Taylor, J. A, Lamb, J. D., & Fujimoto, V. Y. (2011b).
Serum unconjugated bisphenol A concentrations in men may influence embryo quality
indicators during in vitro fertilization. Environmental Toxicology and Pharmacology,
32(2), 319–23. doi:10.1016/j.etap.2011.06.003
Bloom, S. E., Muscarella, D. E., Lee, M. Y., & Rachlinski, M. (1998). Cell death in the avian
blastoderm: resistance to stress-induced apoptosis and expression of anti-apoptotic genes.
Cell Death and Differentiation, 5, 529–538. doi:10.1038/sj.cdd.4400381
Boas, M., Feldt-Rasmussen, U., & Main, K. M. (2012). Thyroid effects of endocrine disrupting
chemicals. Molecular and Cellular Endocrinology, 355(2), 240–8.
doi:10.1016/j.mce.2011.09.005
Braun, J. M., Kalkbrenner, A. E., Calafat, A. M., Yolton, K., Ye, X., Dietrich, K. N., &
Lanphear, B. P. (2011). Impact of early-life bisphenol A exposure on behavior and
executive function in children. Pediatrics, 128(5), 873–82. doi:10.1542/peds.2011-1335
Brevini, T. a L., Cillo, F., Antonini, S., & Gandolfi, F. (2007). Cytoplasmic remodelling and the
acquisition of developmental competence in pig oocytes. Animal Reproduction Science,
98(1-2), 23–38. doi:10.1016/j.anireprosci.2006.10.018
Brevini, T. a L., Lonergan, P., Cillo, F., Francisci, C., Favetta, L. A., Fair, T., & Gandolfi, F.
(2002). Evolution of mRNA polyadenylation between oocyte maturation and first
embryonic cleavage in cattle and its relation with developmental competence. Molecular
Reproduction and Development, 63(4), 510–517. doi:10.1002/mrd.10191
Bromfield, J. J., Coticchio, G., Hutt, K., Sciajno, R., Borini, A., & Albertini, D. F. (2009).
Meiotic spindle dynamics in human oocytes following slow-cooling cryopreservation.
Human Reproduction, 24(9), 2114–2123. doi:10.1093/humrep/dep182
140
Brunet, S., & Verlhac, M. H. (2011). Positioning to get out of meiosis: The asymmetry of
division. Human Reproduction Update, 17(1), 68–75. doi:10.1093/humupd/dmq044
Bushnik, T., Haines, D., Levallois, P., Levesque, J., Van Oostdam, J., & Viau, C. (2010). Lead
and bisphenol A concentrations in the Canadian population. Health Reports, 21, 7–18
Byrne, A. T., Southgate, J., Brison, D. R., & Leese, H. J. (1999). Analysis of apoptosis in the
preimplantation bovine embryo using TUNEL. Journal of Reproduction and Fertility,
117(1), 97–105. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/10645250
Calafat, A. M., Kuklenyik, Z., Reidy, J. a., Caudill, S. P., Ekong, J., & Needham, L. L. (2005).
Urinary concentrations of bisphenol A and 4-Nonylphenol in a human reference
population. Environmental Health Perspectives, 113(4), 391–395. doi:10.1289/ehp.7534
Calafat, A. M., Ye, X., Wong, L.-Y., Reidy, J. A, & Needham, L. L. (2008). Exposure of the
U.S. population to bisphenol A and 4-tertiary-octylphenol: 2003-2004. Environmental
Health Perspectives, 116(1), 39–44. doi:10.1289/ehp.10753
Calder, M. D., Caveney, A. N., Sirard, M. A., & Watson, A. J. (2005). Effect of serum and
cumulus cell expansion on marker gene transcripts in bovine cumulus-oocyte complexes
during maturation in vitro. Fertility and Sterility, 83(4 SUPPL.), 1077–1085.
doi:10.1016/j.fertnstert.2004.12.012
Camargo, L. S. A, Viana, J. H. M., Ramos, a a, Serapião, R. V, de Sa, W. F., Ferreira, A. M, &
do Vale Filho, V. R. (2007). Developmental competence and expression of the Hsp 70.1
gene in oocytes obtained from Bos indicus and Bos taurus dairy cows in a tropical
environment. Theriogenology, 68(4), 626–32. doi:10.1016/j.theriogenology.2007.03.029
Can, A., Semiz, O., & Cinar, O. (2005). Bisphenol-A induces cell cycle delay and alters
centrosome and spindle microtubular organization in oocytes during meiosis. Molecular
Human Reproduction, 11(6), 389–96. doi:10.1093/molehr/gah179
Cantonwine, D., Meeker, J. D., Hu, H., Sánchez, B. N., Lamadrid-Figueroa, H., Mercado-García,
A., Fortenberry, G. Z., Calafat, A. M. & Tellez-Rojo, M. M. (2010). Bisphenol A exposure
in Mexico City and risk of prematurity: a pilot nested case control study. Environmental
Health : A Global Access Science Source, 9, 62. doi:10.1186/1476-069X-9-62
Carabatsos, M. J., Elvin, J., Matzuk, M. M., & Albertini, D. F. (1998). Characterization of oocyte
and follicle development in growth differentiation factor-9-deficient mice. Developmental
Biology, 204(2), 373–384. doi:10.1006/dbio.1998.9087
Carabatsos, M. J., Sellitto, C., Goodenough, D. A, & Albertini, D. F. (2000). Oocyte-granulosa
cell heterologous gap junctions are required for the coordination of nuclear and
cytoplasmic meiotic competence. Developmental Biology, 226, 167–179.
doi:10.1006/dbio.2000.9863
Carwile, J. L., & Michels, K. B. (2011). Urinary bisphenol A and obesity: NHANES 2003-2006.
Environmental Research, 111(6), 825–30. doi:10.1016/j.envres.2011.05.014
141
Caudle, J. R. (2013). Examination of Y chromosome linked gene expression in healthy and
arrested pre-implantation bovine embryos. MSc thesis. The University of Guelph.
Cecconi S, Paro R, Rossi G, Macchiarelli G. (2007). The effects of the endocrine disruptors
dithio- carbamates on the mammalian ovary with particular regard to mancozeb. Current
Pharmaceutical Design, 13, 2989–3004
Cha, K. Y., Barnes, R. B., Marrs, R. P., & Lobo, R. A. (1986). Correlation of the bioactivity of
luteinizing hormone in follicular fluid with oocyte maturity in the spontaneous cycle.
Fertility and Sterility, 45(3), 338-341.
Chapin, R. E., Adams, J., Boekelheide, K., Gray, L. E., Hayward, S. W., Lees, P. S. J., McIntyre,
B. S., Portier, K. M., Schnorr, T. M., Selevan, S. G., Vandenbergh, J. G., & Woskie, S. R.
(2008). NTP-CERHR expert panel report on the reproductive and developmental toxicity
of bisphenol A. Birth Defects Research Part B - Developmental and Reproductive
Toxicology, 83(January), 157–395. doi:10.1002/bdrb.20147
Chen, J., Melton, C., Suh, N., Oh, J. S., Horner, K., Xie, F., Sette, C., Blelloch, R. & Conti, M.
(2011). Genome-wide analysis of translation reveals a critical role for deleted in
azoospermia-like (Dazl) at the oocyte-to-zygote transition. Genes and Development, 25(7),
755–766. doi:10.1101/gad.2028911
Chesnel, F., Wigglesworth, K., & Eppig, J. J. (1994). Acquisition of meiotic competence by
denuded mouse oocytes: Participation of somatic-cell product(s) and cAMP. Dev. Biol.
161, 285–295.
Chian, R.-C., Buckett, W. M., & Tan, S.-L. (2004). In-vitro maturation of human oocytes.
Reproductive BioMedicine Online, 8(2), 148–166. doi:10.1016/S1472-6483(10)60511-1
Chitra, K. C., Latchoumycandane, C., & Mathur, P. P. (2003). Induction of oxidative stress by
bisphenol A in the epididymal sperm of rats. Toxicology, 185, 119–127.
doi:10.1016/S0300-483X(02)00597-8
Choi, J., & Donehower, L. A. (1999). P53 in Embryonic Development: Maintaining a Fine
Balance. Cellular and Molecular Life Sciences : CMLS, 55(1), 38–47. Retrieved from
http://www.ncbi.nlm.nih.gov/pubmed/10065150
Colborn, T., Vom Saal, F. S., & Soto, A. M. (1993). Developmental effects of endocrinedisrupting chemicals in wildlife and humans. Environmental Health Perspectives,
101(July), 378–384. doi:10.1289/ehp.93101378
Collins, J. A., Schandl, C. A., Young, K. K., Vesely, J., & Willingham, M. C. (1997). Major
DNA fragmentation is a late event in apoptosis, 45(7), 923–934.
Combelles, C. M. H., & Albertini, D. F. (2003). Assessment of oocyte quality following repeated
gonadotropin stimulation in the mouse. Biology of Reproduction, 68, 812–821.
doi:10.1095/biolreprod.102.008656
142
Connor, E. E., Wood, D. L., Sonstegard, T. S., Mota, A. F., Bennett, G. L., Williams, J. L., &
Capuco, A. V. (2005). Chromosomal mapping and quantitative analysis of estrogen-related
receptor alpha-1 , estrogen receptors alpha and beta and progesterone receptor in the
bovine mammary gland, 593–603. doi:10.1677/joe.1.06139
Coticchio, G., Guglielmo, M. C., Dal Canto, M., Fadini, R., Mignini Renzini, M., De Ponti, E.,
Brambillasca, F., & Albertini, D. F. (2013). Mechanistic foundations of the metaphase II
spindle of human oocytes matured in vivo and in vitro. Human Reproduction, 28(12),
3271–82. doi:10.1093/humrep/det381
Coticchio, G., Sciajno, R., Hutt, K., Bromfield, J., Borini ,A., Albertini, D.F. (2010).
Comparative analysis of the metaphase II spindle of human oocytes through polarized light
and high-performance confocal microscopy. Fertility and Sterility, 93, 2056–2064.
Craig, Z. R., Wang, W., & Flaws, J. A. (2011). Endocrine-disrupting chemicals in ovarian
function: effects on steroidogenesis, metabolism and nuclear receptor signaling.
Reproduction, 142, 633–646. doi:10.1530/REP-11-0136
Csanády, G. A., Oberste-Frielinghaus, H. R., Semder, B., Baur, C., Schneider, K. T., Filser, J. G.
(2002). Distribution and unspecific protein binding of the xenoestrogens bisphenol A and
daidzein. Archives of Toxicology, 76, 299–305.
Cui, X.-S., Li, X.-Y., Yin, X.-J., Kong, I. K., Kang, J.-J., & Kim, N.-H. (2007). Maternal gene
transcription in mouse oocytes: genes implicated in oocyte maturation and fertilization.
The Journal of Reproduction and Development, 53(2), 405–418. doi:10.1262/jrd.18113
Da Broi, M. G., Malvezzi, H., Paz, C. C. P., Ferriani, R. a, & Navarro, P. A. A. S. (2014).
Follicular fluid from infertile women with mild endometriosis may compromise the
meiotic spindles of bovine metaphase II oocytes. Human Reproduction (Oxford, England),
29(2), 315–23. doi:10.1093/humrep/det378
Darras, V. M., Van Herck, S. L. J., Heijlen, M., & De Groef, B. (2011). Thyroid hormone
receptors in two model species for vertebrate embryonic development: chicken and
zebrafish. Journal of Thyroid Research, 2011, 402320. doi:10.4061/2011/402320
Darzynkiewicz, Z., Bedner E., Traganos, F. (2001). Difficulties and pitfalls in analysis of
apoptosis. Methods in Cell Biology, 63, 527-546.
De La Fuente, R., & Eppig, J. J. (2001). Transcriptional activity of the mouse oocyte genome:
companion granulosa cells modulate transcription and chromatin remodeling.
Developmental Biology, 229, 224–236. doi:10.1006/dbio.2000.9947
De La Fuente, R., Viveiros, M. M., Burns, K. H., Adashi, E. Y., Matzuk, M. M., & Eppig, J. J.
(2004). Major chromatin remodeling in the germinal vesicle (GV) of mammalian oocytes
is dispensable for global transcriptional silencing but required for centromeric
heterochromatin function. Developmental Biology, 275(2), 447–458.
doi:10.1016/j.ydbio.2004.08.028
143
De Sousa, P. A., Watson, A. J., Schultz, G. A., & Bilodeau-Goeseels, S. (1998). Oogenetic and
zygotic gene expression directing early bovine embryogenesis: A review. Molecular
Reproduction and Development, 51(1), 112–121. doi:10.1002/(SICI)10982795(199809)51:1<112::AID-MRD14>3.0.CO;2-9
Dean, J. H., Chapman, M. G., & Sullivan, E. A. (2010). The effect on human sex ratio at birth by
assisted reproductive technology (ART) procedures - An assessment of babies born
following single embryo transfers, Australia and New Zealand, 2002-2006. BJOG: An
International Journal of Obstetrics and Gynaecology, 117, 1628–1634. doi:10.1111/j.14710528.2010.02731.x
Dennery, P. A. (2007). Effects of oxidative stress on embryonic development. Birth Defects
Research Part C: Embryo Today, 81(3), 155-162.
Di Pasquale, E., Beck-Peccoz, P., & Persani, L. (2004). Hypergonadotropic ovarian failure
associated with an inherited mutation of human bone morphogenetic protein-15 (BMP15)
gene. American Journal of Human Genetics, 75, 106–111. doi:10.1086/422103
Diaz, F. J., Brien, M. J., Wigglesworth, K., & Eppig, J. J. (2006). The preantral granulosa cell to
cumulus cell transition in the mouse ovary: Development of competence to undergo
expansion. Developmental Biology, 299, 91–104. doi:10.1016/j.ydbio.2006.07.012
Diaz, F. J., Wigglesworth, K., & Eppig, J. J. (2007). Oocytes determine cumulus cell lineage in
mouse ovarian follicles. Journal of Cell Science, 120, 1330–1340. doi:10.1242/jcs.000968
Dobrzynska, M. M. & Radzikowska J. (2013). Genotoxicity and reproductive toxicity of
bisphenol A and X-ray/bisphenol A combination in male mice. Drug and Chemical
Toxicology, 36, 19–26.
Dodds, E. C., & Lawson W. (1936). Synthetic estrogenic agents without the phenanthrene
nucleus. Nature, 137(996).
Doerge, D. R., Twaddle, N. C., Vanlandingham, M., & Fisher, J. W. (2011). Pharmacokinetics of
Bisphenol A in neonatal and adult CD-1 mice: Inter-species comparisons with SpragueDawley rats and rhesus monkeys. Toxicology Letters, 207(3), 298–305.
doi:10.1016/j.toxlet.2011.09.020
Dolinoy, D. C., Huang, D., & Jirtle, R. L. (2007). Maternal nutrient supplementation counteracts
bisphenol A-induced DNA hypomethylation in early development. Proceedings of the
National Academy of Sciences of the United States of America, 104(32), 13056–61.
doi:10.1073/pnas.0703739104
Donnison, M., & Pfeffer, P. L. (2004). Isolation of genes associated with developmentally
competent bovine oocytes and quantitation of their levels during development. Biology of
Reproduction, 71(6), 1813–1821. doi:10.1095/biolreprod.104.032367
144
Dunphy, W. G., Brizuela, L., Beach, D., & Newport, J. (1988). The Xenopus cdc2 protein is a
component of MPF, a cytoplasmic regulator of mitosis. Cell, 54(3), 423–431.
doi:10.1016/0092-8674(88)90205-X
Duranthon, V., Watson, A. J., & Lonergan, P. (2008). Preimplantation embryo programming:
transcription, epigenetics, and culture environment. Reproduction, 135(2), 141–50.
doi:10.1530/REP-07-0324
Ehrlich, S., Williams, P. L., Missmer, S. A., Flaws, J. A., Berry, K. F., Calafat, A. M., Ye, X.,
Petrozza, J. C., Wright, D., & Hauser, R. (2012a). Urinary bisphenol A Concentrations and
implantation failure among women undergoing in vitro fertilization. Environmental Health
Perspectives, 120, 978-983.
Ehrlich, S., Willliams, P. L., Missmer, S. A., Flaws, J. A., Ye, X., Calafat, A. M., Petrozza, J. C.,
Wright, D., & Haiser, R. (2012b). Urinary bisphenol A concentrations and early
reproductive health outcomes among women undergoing IVF. Human Reproduction, 27,
3583-3359.
Eichenlaub-Ritter, U., Shen, Y., & Tinneberg, H.-R. (2002). Manipulation of the oocyte: possible
damage to the spindle apparatus. Reproductive Biomedicine Online, 5(2), 117–124.
doi:10.1016/S1472-6483(10)61613-6
Eichenlaub-Ritter, U., Vogt, E., Cukurcam, S., Sun, F., Pacchierotti, F., & Parry, J. (2008).
Exposure of mouse oocytes to bisphenol A causes meiotic arrest but not aneuploidy.
Mutation Research, 651(1-2), 82–92. doi:10.1016/j.mrgentox.2007.10.014
El Mouatassim, S., Guérin, P., & Ménézo, Y. (1999). Expression of genes encoding antioxidant
enzymes in human and mouse oocytes during the final stages of maturation. Molecular
Human Reproduction, 5(8), 720–725.
El-Sayed, A., Hoelker, M., Rings, F., Salilew, D., Jennen, D., Tholen, E., Sirard, M-A.,
Schellander, K., & Tesfaye, D. (2006). Large-scale transcriptional analysis of bovine
embryo biopsies in relation to pregnancy success after transfer to recipients. Physiological
Genomics, 28, 84–96. doi:10.1152/physiolgenomics.00111.2006
Elsby, R., Maggs, J. L., Ashby, J., & Park, B. K. (2001). Comparison of the modulatory effects
of human and rat liver microsomal metabolism on the estrogenicity of bisphenol A:
implications for extrapolation to humans. The Journal of Pharmacology and Experimental
Therapeutics, 297(1), 103–113.
Eppig, J. J. (2001). Oocyte control of ovarian follicular development and function in mammals.
Reproduction, 122, 829–838. doi:10.1530/reprod/122.6.829
Eppig, J. J., O’Brien, M., & Wigglesworth, K. (1996). Mammalian oocyte growth and
development in vitro. Molecular Reproduction and Development, 44, 260–273.
doi:10.1002/(SICI)1098-2795(199606)44:2<260::AID-MRD17>3.0.CO;2-6
145
Eppig, J. J., Wigglesworth, K., & Pendola, F. L. (2002). The mammalian oocyte orchestrates the
rate of ovarian follicular development. Proceedings of the National Academy of Sciences
of the United States of America, 99(5), 2890–2894. doi:10.1073/pnas.052658699
Ertzeid, G., & Storeng, R. (2001). The impact of ovarian stimulation on implantation and fetal
development in mice. Human Reproduction, 16(2), 221–225. doi:10.1093/humrep/16.2.221
Evsikov, A. V, Graber, J. H., Brockman, J. M., Hampl, A., Holbrook, A. E., Singh, P., Eppig, J.
J., Solter, D. & Knowles, B. B (2006). Cracking the egg : molecular dynamics and
evolutionary aspects of the transition from the fully grown oocyte to embryo. Genes &
Development, 20, 2713-2727.
Farrell P. B. & Foote R. H. (1995). Beneficial effects of culturing rabbit zygotes to blastocysts in
5% oxygen and 10% carbon dioxide. Journal of Reproduction and Fertility, 103(127–130).
Favetta, L. A, Madan, P., Mastromonaco, G. F., St John, E. J., King, W. A., & Betts, D. H.
(2007). The oxidative stress adaptor p66Shc is required for permanent embryo arrest in
vitro. BMC Developmental Biology, 7, 132. doi:10.1186/1471-213X-7-132
Favetta, L. A., Robert, C., St. John, E. J., Betts, D. H., & King, W. A. (2004). P66Shc, but not
P53, is involved in early arrest of in vitro-produced bovine embryos. Molecular Human
Reproduction, 10(6), 383–392. doi:10.1093/molehr/gah057
First, N. L., Leibfried-Rutledge, M.L., & Sirard, M.A. (1988). Cytoplasmic control of oocyte
maturation and species differences in the development of maturational competence.
Progress in Clinical Biological Research, 267, 1–46.
Fowler, P. a., Bellingham, M., Sinclair, K. D., Evans, N. P., Pocar, P., Fischer, B., Schaedlich,
K., Schmidt, J-S., Amegaza, M. R., Bhattacharya, S., Rhind, S. M., & O’Shaughnessy, P. J.
(2012). Impact of endocrine-disrupting compounds (EDCs) on female reproductive health.
Molecular and Cellular Endocrinology, 355(2), 231–239. doi:10.1016/j.mce.2011.10.021.
Fujimoto, V. Y., Kim, D., vom Saal, F. S., Lamb, J. D., Taylor, J. A., & Bloom, M. S. (2011).
Serum unconjugated bisphenol A concentrations in women may adversely influence oocyte
quality during in vitro fertilization. Fertility and Sterility, 95(5), 1816–1819.
doi:10.1016/j.fertnstert.2010.11.008.
Fukuda, M., Fukuda, K., Shimizu, T., & Møller, H. (1998). Decline in sex ratio at birth after
Kobe earthquake. Human Reproduction (Oxford, England), 13(8), 2321–2322.
Gandolfi, F., Pocar, P., Brevini, T. a L., & Fischer, B. (2002). Impact of endocrine disrupters on
ovarian function and embryonic development. Domestic Animal Endocrinology, 23(1-2),
189–201. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/12142237
Garrido, N., Navarro, J., Remohí, J., Simón, C., & Pellicer, A. (2000). Follicular hormonal
environment and embryo quality in women with endometriosis. Human Reproduction
Update, 6(1), 67–74.
146
Gautier, J., Minshull, J., Lohka, M., Glotzer, M., Hunt, T., & Maller, J. L. (1990). Cyclin is a
component of maturation-promoting factor from Xenopus. Cell, 60(3), 487–494.
doi:10.1016/0092-8674(90)90599-A
Gautier, J., Norbury, C., Lohka, M., Nurse, P., & Maller, J. (1988). Purified maturationpromoting factor contains the product of a Xenopus homolog of the fission yeast cell cycle
control gene cdc2+. Cell, 54(3), 433–439. doi:10.1016/0092-8674(88)90206-1
Gendelman, M., & Roth, Z. (2012). Seasonal effect on germinal vesicle-stage bovine oocytes is
further expressed by alterations in transcript levels in the developing embryos associated
with reduced developmental competence. Biology of Reproduction, 86(1), 1–9.
doi:10.1095/biolreprod.111.092882
Ghisari, M., & Bonefeld-Jorgensen, E. C. (2005). Impact of environmental chemicals on the
thyroid hormone function in pituitary rat GH3 cells. Molecular and Cellular
Endocrinology, 244, 31–41. doi:10.1016/j.mce.2005.01.013
Gilchrist, R. B., Lane, M., & Thompson, J. G. (2008). Oocyte-secreted factors: Regulators of
cumulus cell function and oocyte quality. Human Reproduction Update, 14(2), 159–177.
doi:10.1093/humupd/dmm040
Gjørret, J. O., Knijn, H. M., Dieleman, S. J., Avery, B., Larsson, L., & Maddox-hyttel, P. (2003).
Chronology of apoptosis in bovine embryos produced in vivo and in vitro, 1200(May),
1193–1200. doi:10.1095/biolreprod.102.013243
Gronemeyer, H., Gustafsson, J.-A., & Laudet, V. (2004). Principles for modulation of the
nuclear receptor superfamily. Nature Reviews. Drug Discovery, 3(November), 950–964.
doi:10.1038/nrd1551
Guérin, P., El Mouatassim, S., & Ménézo, Y. (2001). Oxidative stress and protection against
reactive oxygen species in the pre-implantation embryo and its surroundings. Human
Reproduction Update, 7(2), 175–189. doi:10.1093/humupd/7.2.175
Guglielmino, M. R., Santonocito, M., Vento, M., Ragusa, M., Barbagallo, D., Borzì, P.,
Casciano, I., Banelli, B., Barbieri, O., Astigiano, S., Scollo, P., Romani, M., Purrello, N., &
Di Pietro, C. (2011). TAp73 is downregulated in oocytes from women of advanced
reproductive age. Cell Cycle, 10(19), 3253–3256. doi:10.4161/cc.10.19.17585
Guo, R., Ebenezer, V., & Ki, J. S. (2012). Transcriptional responses of heat shock protein 70
(Hsp70) to thermal, bisphenol A, and copper stresses in the dinoflagellate Prorocentrum
minimum. Chemosphere, 89, 512–520. doi:10.1016/j.chemosphere.2012.05.014
Gutiérrez-Adán, A., Oter M., Martinez-Madrid B, Pintado B. & De La Fuente J. (2000).
Differential expression of two genes located on the X chromosome between male and
female in vitro-produced bovine embryos at the blastocyst stage. Molecular Reproduction
and Development, 55, 146–151.
147
Gutiérrez-Adán, A, Lonergan, P., Rizos, D., Ward, F. A, Boland, M. P., Pintado, B., & De La, F.
J. (2001). Effect of the in vitro culture system on the kinetics of blastocyst development
and sex ratio of bovine embryos. Theriogenology, 55, 1117–1126.
Gutiérrez-Adán, a, Perez-Crespo, M., Fernandez-Gonzalez, R., Ramirez, M. A., Moreira, P.,
Pintado, B., Lonergan, P., & Rizos, D. (2006). Developmental consequences of sexual
dimorphism during pre-implantation embryonic development. Reproduction in Domestic
Animals, 41 (Suppl. 2), 54-62. doi:10.1111/j.1439-0531.2006.00769.x
Hamdoun, A., & Epel, D. (2007). Changing world protection strategies in early development : a
general strategy, 104(6), 1745–1750.
Hamilton, C. K., A. Combe, J. Caudle, F. A. Ashkar, A. D. Macaulay, P. Blondin, and W. A.
King. 2012. A novel approach to sexing bovine blastocysts using male-specific gene
expression. Theriogenology, 77(8), 1587-96
Hamilton, C. K., Revay, T., Domander, R., Favetta, L. A., & King, W. A. (2011). A large
expansion of the HSFY gene family in cattle shows dispersion across Yq and testis-specific
expression. PLoS ONE, 6(3). doi:10.1371/journal.pone.0017790
Hansen, P. J., & Fear, J. M. (2011). Cheating death at the dawn of life: Developmental control of
apoptotic repression in the preimplantation embryo. Biochemical and Biophysical
Research Communications, 413(2), 155–158. doi:10.1016/j.bbrc.2011.08.064
Hardy, K. (1999). Apoptosis in the human embryo. Reviews of Reproduction, 4(3), 125–134.
doi:10.1530/ror.0.0040125
Hardy, K., Handyside, A. H., & Winston R, M. (1989). The human blastocyst: Cell number,
death, and allocation during late preimplantation development in vitro. Development, 107,
597–604.
Harvey, a J., Kind, K. L., Pantaleon, M., Armstrong, D. T., & Thompson, J. G. (2004). Oxygenregulated gene expression in bovine blastocysts. Biology of Reproduction, 71, 1108–1119.
doi:10.1095/biolreprod.104.028639
Harvey, M.B., Arcellana-Panlilio, M.Y., Zhang, X. Schultz, G A., & Watson, A. J. (1995).
Expression of genes encoding antioxidant enzymes in preimplantation mouse and cow
embryos and primary bovine oviduct cultures employed for embryo coculture. Biology of
Reproduction, 53, 532-540.
Hassold, T., & Hunt, P. (2001). To err (meiotically) is human: the genesis of human aneuploidy.
Nature Reviews. Genetics, 2, 280–291. doi:10.1038/35066065
Heimeier, R. a, Das, B., Buchholz, D. R., & Shi, Y.-B. (2009). The xenoestrogen bisphenol A
inhibits postembryonic vertebrate development by antagonizing gene regulation by thyroid
hormone. Endocrinology, 150(6), 2964–73. doi:10.1210/en.2008-1503
148
Hendrey, J., & Kola, I. (1991). Thermolability of mouse oocytes is due to the lack of expression
and/or inducibility of Hsp70. Molecular Reproduction and Development, 28, 1–8.
doi:10.1002/mrd.1080280102
Hiroi, H., Momoeda, M., Inoue, S., Tsuchiya, F., Matsumi, H., Tsutsumi, O., Muramatsu, M., &
Taketani, Y. (1999). Stage-specific expression of estrogen receptor subtypes and estrogen
responsive finger protein in preimplantational mouse embryos. Endocrine Journal, 46(1),
153–158. doi:10.1507/endocrj.46.153
Hodges, C. A., Ilagan, A., Jennings, D., Keri, R., Nilson, J., & Hunt, P. A. (2002). Experimental
evidence that changes in oocyte growth influence meiotic chromosome segregation, 17(5),
1171–1180.
Hodgman, R., Tay, J., Mendez, R., & Richter, J. D. (2001). CPEB phosphorylation and
cytoplasmic polyadenylation are catalyzed by the kinase IAK1/Eg2 in maturing mouse
oocytes. Development, 128(14), 2815–2822.
Hou, Q., & Gorski, J. (1993). Estrogen receptor and progesterone receptor genes are expressed
differentially in mouse embryos during preimplantation development. Proceedings of the
National Academy of Sciences of the United States of America, 90, 9460–9464.
doi:10.1073/pnas.90.20.9460
Hou, Q., Paria, B. C., Mui, C., Dey, S. K., & Gorski, J. (1996). Immunolocalization of estrogen
receptor protein in the mouse blastocyst during normal and delayed implantation.
Proceedings of the National Academy of Sciences of the United States of America, 93,
2376–2381. doi:10.1073/pnas.93.6.2376
Howdeshell, K. L., Hotchkiss, A. K., Thayer, K. A, Vandenbergh, J. G., & vom Saal, F. S.
(1999). Exposure to bisphenol A advances puberty. Nature, 401, 763–764.
doi:10.1038/44517
Hu, F. B., Manson, J. E., Stampfer, M. J., Colditz, G., Liu, S., Solomon, C. G., & Willett, W. C.
(2001). Diet, lifestyle, and the risk of type 2 diabetes mellitus in women. New England
Journal of Medicine, 345, 790–797.
Hu, W., Zheng, T., & Wang, J. (2011). Regulation of Fertility by the p53 Family Members.
Genes & Cancer, 2(4), 420–430. doi:10.1177/1947601911408892
Huarte, J., Belin, D., Vassalli, A., Strickland, S., & Vassalli, J. D. (1987). Meiotic maturation of
mouse oocytes triggers the translation and polyadenylation of dormant tissue-type
plasminogen activator mRNA. Genes & Development, 1(10), 1201–1211.
doi:10.1101/gad.1.10.1201
Hunt, P. A, & Hassold, T. J. (2008). Human female meiosis: what makes a good egg go bad?
Trends in Genetics : TIG, 24(2), 86–93. doi:10.1016/j.tig.2007.11.010
149
Hunt, P. A, Koehler, K. E., Susiarjo, M., Hodges, C. A, Ilagan, A., Voigt, R. C., Thomas, S.,
Thomas, B. F., & Hassold, T. J. (2003). Bisphenol A exposure causes meiotic aneuploidy
in the female mouse. Current Biology, 13(7), 546–53. doi:10.1016/S
Hunter, A. G. & Moor, R. M. (1987). Stage-dependent effects of inhibiting ribonucleic acids and
protein synthesis on meiotic maturation of bovine oocytes in vitro. Journal of Dairy
Science, 70, 1646–1651
Hyttel, P., Callesen, H., Greve, T. (1989). A comparative ultrastructural study of in vivo versus
in vitro fertilization of bovine oocytes. Anatomy and Embryology, 179, 435-442.
Ibáñez, E., Sanfins, A., Combelles, C. M. H., Overström, E. W., & Albertini, D. F. (2005).
Genetic strain variations in the metaphase-II phenotype of mouse oocytes matured in vivo
or in vitro. Reproduction, 130, 845–855. doi:10.1530/rep.1.00558
Ikezuki, Y., Tsutsumi, O., Takai, Y., Kamei, Y., & Taketani, Y. (2002). Determination of
bisphenol A concentrations in human biological fluids reveals significant early prenatal
exposure. Human Reproduction, 17(11), 2839–41. Retrieved from
http://www.ncbi.nlm.nih.gov/pubmed/12407035
Ishibashi, H., Watanabe, N., Matsumura, N., Hirano, M., Nagao, Y., Shiratsuchi, H., Kohra, S.,
Yoshihara, S., & Arizono, K. (2005). Toxicity to early life stages and an estrogenic effect
of a bisphenol A metabolite, 4-methyl-2,4-bis(4-hydroxyphenyl)pent-1-ene on the medaka
(Oryzias latipes). Life Sciences, 77, 2643–2655. doi:10.1016/j.lfs.2005.03.025
Ishihara, K., Warita, K., Tanida, T., Sugawara, T., Kitagawa, H., & Hoshi, N. (2007). Does
paternal exposure to 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) affect the sex ratio of
offspring? The Journal of Veterinary Medical Science / the Japanese Society of Veterinary
Science, 69, 347–352. doi:10.1292/jvms.69.347
Iwamuro, S., Sakakibara, M., Terao, M., Ozawa, A., Kurobe, C., Shigeura, T., Kato, M., &
Kikuyama, S. (2003). Teratogenic and anti-metamorphic effects of bisphenol A on
embryonic and larval Xenopus laevis. General and Comparative Endocrinology, 133(2),
189–198. doi:10.1016/S0016-6480(03)00188-6
Iwata, H., Shiono, H., Kon, Y., Matsubara, K., Kimura, K., Kuwayama, T., & Monji, Y. (2008).
Effects of modification of in vitro fertilization techniques on the sex ratio of the resultant
bovine embryos. Animal Reproduction Science, 105(3-4), 234–44.
doi:10.1016/j.anireprosci.2007.03.006
Jacobson, M. D., Weil, M., & Raff, M. C. (1997). Programmed cell death in animal
development. Cell, 88(3), 347–354. doi:9039261
Jakubiczka, S., Schnieders, F., & Schmidtke, J. (1993). A bovine homologue of the human TSPY
gene. Genomics, 17, 732–735.
James, W. H. (1998). Was the widespread decline in sex ratios at birth caused by reproductive
hazards? Human Reproduction, 13(4), 1083–1084. doi:10.1093/humrep/13.4.1083
150
Jiménez, A., Madrid-Bury, N., Fernandez, R., Pérez-Garnelo, S., Moreira, P., Pintado, B., …
Gutiérrez-Adan, A. (2003). Hyperglycemia-induced apoptosis affects sex ratio of bovine
and murine preimplantation embryos. Molecular Reproduction and Development, 65, 180–
187. doi:10.1002/mrd.10286
Jin, P., Wang, X., Chang, F., Bai, Y., Li, Y., Zhou, R., & Chen, L. (2013). Low dose bisphenol A
impairs spermatogenesis by suppress- ing reproductive hormone production and promoting
germ cell apoptosis in adult rats. Journal of Biomedical Research, 27, 135–144.
Johnson, M. H., Nasr-Esfahani, M. H. (1994). Radical solutions and culture problems: could free
oxygen radicals be responsible for the impaired development of preimplantation
mammalian embryos in vitro? Bioessays, 16, 31-38.
Jones, G. M., Cram, D. S., Song, B., Magli, M. C., Gianaroli, L., Lacham-Kaplan, O., Findlay, J.
K., Jenkin, G., & Trounson, A. O. (2008). Gene expression profiling of human oocytes
following in vivo or in vitro maturation. Human Reproduction, 23(5), 1138–1144.
doi:10.1093/humrep/den085
Jurisicova, A., Latham, K., Casper, R. & Varmuza, S. (1998) Expression and regulation of genes
associated with cell death during murine preimplantation embryo development. Molecular
Reproduction and Development, 51, 243-253.
Kabuto, H., Hasuike, S., Minagawa, N., & Shishibori, T. (2003). Effects of bisphenol A on the
metabolisms of active oxygen species in mouse tissues. Environmental Research, 93, 31–
35. doi:10.1016/S0013-9351(03)00062-8
Källén, B., Finnström, O., Nygren, K. G., & Olausson, P. O. (2005). In vitro fertilization (IVF) in
Sweden: Infant outcome after different IVF fertilization methods. Fertility and Sterility,
84(3), 611–617. doi:10.1016/j.fertnstert.2005.02.038
Kang, J.-H., Kondo, F., & Katayama, Y. (2006). Human exposure to bisphenol A. Toxicology,
226(2-3), 79–89. doi:10.1016/j.tox.2006.06.009
Kawarsky, S. J., & King, W. a. (2001). Expression and localisation of heat shock protein 70 in
cultured bovine oocytes and embryos. Zygote, 9(1), 39–50. Retrieved from
http://www.ncbi.nlm.nih.gov/pubmed/11273032
Kidane, D., Sakkas, D., Nottoli, T., McGrath, J., & Sweasy, J. B. (2013). Kinesin 5B (KIF5B) is
required for progression through female meiosis and proper chromosomal segregation in
mitotic cells. PLoS ONE, 8(4). doi:10.1371/journal.pone.0058585
Kinyamu, H. K., & Archer, T. K. (2003). Estrogen receptor-dependent proteasomal degradation
of the glucocorticoid receptor is coupled to an increase in mdm2 protein expression.
Molecular and Cellular Biology, 23(16), 5867–5881. doi:10.1128/MCB.23.16.58675881.2003
Ko, L. J., & Prives, C. (1996). p53: Puzzle and paradigm. Genes and Development, 10, 1054–
1072. doi:10.1101/gad.10.9.1054
151
Kobayashi, S., Isotani, A., Mise, N., Yamamoto, M., Fujihara, Y., Kaseda, K., Nakanishi, T.,
Ikawa, M., Hamada, H., Abe, K., & Okabe, M. (2006). Comparison of gene expression in
male and female mouse blastocysts revealed imprinting of the X-linked gene, Rhox5/Pem,
at preimplantation stages. Current Biology, 16, 166–172. doi:10.1016/j.cub.2005.11.071
Krackow, S. (1997). Effects of mating dynamics and crowding on sex ratio variance in mice.
Journal of Reproduction and Fertility, 110, 87–90. doi:10.1530/jrf.0.1100087
Kregel, K. C. (2002). Heat shock proteins: modifying factors in physiological stress responses
and acquired thermotolerance. Journal of Applied Physiology (Bethesda, Md. : 1985), 92,
2177–2186. doi:10.1152/japplphysiol.01267.2001
Kreiner D., Liu, H. C., Itskovitz, J., Veeck, L., & Rosenwaks, Z. (1987). Follicular fluid estradiol
and progesterone are markers of preovulatory oocyte quality. Fertility and Sterility, 48
(991–994).
Krisher, R. L. (2013). In vivo and in vitro environmental effects on mammalian oocyte quality.
Annual Review of Animal Biosciences, 1(1), 393–417. doi:10.1146/annurev-animal031412-103647
Krisher, R. L., & Bavister, B. D. (1999). Enhanced glycolysis after maturation of bovine oocytes
in vitro is associated with increased developmental competence. Molecular Reproduction
and Development, 53, 19–26. doi:10.1002/(SICI)1098-2795(199905)53:1<19::AIDMRD3>3.0.CO;2-U
Kwintkiewicz, J., Nishi, Y., Yanase, T., & Giudice, L. C. (2010). Peroxisome proliferatoractivated receptor-gamma mediates bisphenol A inhibition of FSH-stimulated IGF-1,
aromatase, and estradiol in human granulosa cells. Environmental Health Perspectives,
118(3), 400–6. doi:10.1289/ehp.0901161
Lahnsteiner, F., Berger, B., Kletzl, M., & Weismann, T. (2005). Effect of bisphenol A on
maturation and quality of semen and eggs in the brown trout, Salmo trutta f. fario. Aquatic
Toxicology, 75, 213–224.
Lane, E. A, & Hyde, T. S. (1973). Effect of maternal stress on fertility and sex ratio: a pilot study
with rats. Journal of Abnormal Psychology, 82(1), 78–80.
Lane, S. I. R., Chang, H. Y., Jennings, P. C., & Jones, K. T. (2010). The Aurora kinase inhibitor
ZM447439 accelerates first meiosis in mouse oocytes by overriding the spindle assembly
checkpoint. Reproduction, 140(4), 521–530. doi:10.1530/REP-10-0223
Larson, M. a, Kimura, K., Kubisch, H. M., & Roberts, R. M. (2001). Sexual dimorphism among
bovine embryos in their ability to make the transition to expanded blastocyst and in the
expression of the signaling molecule IFN-tau. Proceedings of the National Academy of
Sciences of the United States of America, 98(17), 9677–9682.
doi:10.1073/pnas.171305398
152
Laurent, P., Elduque, C., Hayes, H., Saunier, K., Eggen, A, & Levéziel, H. (2000). Assignment
of 60 human ESTs in cattle. Mammalian Genome, 11(9), 748–754.
doi:10.1007/s003350010159
Lawson, C., Gieske, M., Murdoch, B., Ye, P., Li, Y., Hassold, T., & Hunt, P. A. (2011). Gene
expression in the fetal mouse ovary is altered by exposure to low doses of bisphenol A.
Biology of Reproduction, 84(1), 79–86. doi:10.1095/biolreprod.110.084814
Lazzari, G., Wrenzycki, C., Herrmann, D., Duchi, R., Kruip, T., Niemann, H., & Galli, C.
(2002). Cellular and molecular deviations in bovine in vitro-produced embryos are related
to the large offspring syndrome. Biology of Reproduction, 67, 767–775.
doi:10.1095/biolreprod.102.004481
Lee, K.-B., Folger, J. K., Rajput, S. K., & Smith, G. W. (2014b). Temporal regulation of mRNAs
for select bone morphogenetic proteins (BMP), BMP receptors and their associated SMAD
proteins during bovine early embryonic development: effects of exogenous BMP2 on
embryo developmental progression. Reproductive Biology and Endocrinology, 12(1), 67.
doi:10.1186/1477-7827-12-67
Lee, S., Kim, E., & Hyun, S-H. (2014). Toxicity evaluation of ethanol treatment during in vitro
maturation of porcine oocytes and subsequent embryonic development following
parthenogenetic activation and in vitro fertilization. International Journal of Molecular
Medicine. 34, 1372-1380.
Leist, M. & Jaattela, M. (2001). Four deaths and a funeral: from caspases to alternative
mechanisms. Nature Reviews Molecular Cell Biology, 2, 589–598.
Lenie, S., Cortvrindt, R., Eichenlaub-Ritter, U., & Smitz, J. (2008). Continuous exposure to
bisphenol A during in vitro follicular development induces meiotic abnormalities. Mutation
Research, 651(1-2), 71–81. doi:10.1016/j.mrgentox.2007.10.017
Lequarre, a S., Grisart, B., Moreau, B., Schuurbiers, N., Massip, A, & Dessy, F. (1997). Glucose
metabolism during bovine preimplantation development: analysis of gene expression in
single oocytes and embryos. Mol Reprod Dev, 48(April), 216–226. Retrieved from
http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?cmd=Retrieve&db=PubMed&dopt=Citatio
n&list_uids=9291471
Lequarre, A. S., Traverso, J. M., Marchandise, J., & Donnay, I. (2004). Poly(A) RNA is reduced
by half during bovine oocyte maturation but increases when meiotic arrest is maintained
with CDK inhibitors. Biology of Reproduction, 71(2), 425–431.
doi:10.1095/biolreprod.103.026724
Leroy, J. L. M. R., Valckx, S., Sturmey, R., Bossaert, P., Hoeck, V. Van, & Bols, P. E. J. (2012).
Maternal metabolic health and oocyte quality : the role of the intrafollicular environment.
Animal Reproduction, 9, 777–788.
Leroy, J. L. M. R., Vanholder, T., Opsomer, G., Van Soom, a, & de Kruif, A. (2006). The in
vitro development of bovine oocytes after maturation in glucose and beta-hydroxybutyrate
153
concentrations associated with negative energy balance in dairy cows. Reproduction in
Domestic Animals, 41, 119–123. doi:10.1111/j.1439-0531.2006.00650.x
Levy, G., Lutz, I., Kruger, A., & Kloas, W. (2004). Bisphenol A induces feminization in
Xenopus laevis tadpoles. Environmental Research, 94, 102-111.
Li, A., Chandrakanthan, V., Chami, O., Lozano, G., & O'Neill, C. (2005). Culture in vitro
increases p53 expression in preimplantation embryos which compromises their
developmental competence, pp. 1-16, Australia (abstract SSR 2005).
Li, D. K., Zhou, Z., Miao, M., He, Y., Wang, J., Ferber, J., Herrinton, L. J., Gao, E., & Yuan, W.
(2011). Urine bisphenol-A (BPA) level in relation to semen quality. Fertility and Sterility,
95(2), 625-630. doi: 10.1016/j.fertnstert.2010.09.026.
Li, R., & Albertini, D. F. (2013). The road to maturation: somatic cell interaction and selforganization of the mammalian oocyte. Nature Reviews. Molecular Cell Biology, 14(3),
141–52. doi:10.1038/nrm3531
Li, Y., Feng, H. L., Cao, Y. J., Zheng, G. J., Yang, Y., Mullen, S., Critser, J. K., & Chen, Z. J.
(2006). Confocal microscopic analysis of the spindle and chromosome configurations of
human oocytes matured in vitro. Fertility and Sterility, 85(4), 827–832.
doi:10.1016/j.fertnstert.2005.06.064
Lichnovský, V., Kolár, Z., Murray, P., Hlobilková, a, Cernochová, D., Pospísilová, E., &
Vojtesek, B., & Nenutil, R. (1998). Differences in p53 and Bcl-2 expression in relation to
cell proliferation during the development of human embryos. Molecular Pathology, 51,
131–137.
Liu, L., Trimarchi, J. R., Navarro, P., Blasco, M. a., & Keefe, D. L. (2003). Oxidative stress
contributes to arsenic-induced telomere attrition, chromosome instability, and apoptosis.
Journal of Biological Chemistry, 278, 31998–32004. doi:10.1074/jbc.M303553200
Liu, R., Xing, L., Kong, D., Jiang, J., Shang, L., & Hao, W. (2013). Bisphenol A inhibits
proliferation and induces apoptosis in micromass cultures of rat embryonic midbrain cells
through the JNK, CREB and p53 signaling pathways. Food and Chemical Toxicology, 52,
76–82. doi:10.1016/j.fct.2012.10.033
Lonergan, P., Gutiérrez-Adán, A., Rizos, D., Pintado, B., La Fuente, J. De, & Boland, M. P.
(2003a). Relative messenger RNA abundance in bovine oocytes collected in vitro or in
vivo before and 20 hr after the preovulatory luteinizing hormone surge. Molecular
Reproduction and Development, 66, 297–305. doi:10.1002/mrd.10357
Lonergan, P., Rizos, D., Gutierrez-Adán, A., Moreira, P. M., Pintado, B., de la Fuente, J., &
Boland, M. P. (2003b). Temporal divergence in the pattern of messenger RNA expression
in bovine embryos cultured from the zygote to blastocyst stage in vitro or in vivo. Biology
of Reproduction, 69(4), 1424–1431. doi:10.1095/biolreprod.103.018168
154
Luna, M., Duke, M., Copperman, A., Grunfeld, L., Sandler, B., & Barritt, J. (2007). Blastocyst
embryo transfer is associated with a sex-ratio imbalance in favor of male offspring.
Fertility and Sterility, 87(3), 519–23. doi:10.1016/j.fertnstert.2006.06.058
Luzzo, K. M., Wang, Q., Purcell, S. H., Chi, M., Jimenez, P. T., Grindler, N., Schedl T., &
Moley, K. H. (2012). High fat diet induced developmental defects in the mouse: oocyte
meiotic aneuploidy and fetal growth retardation/brain defects. PloS One, 7(11), e49217.
doi:10.1371/journal.pone.0049217
Macaulay, a D., Hamilton, C. K., Bartlewski, P. M., & King, W. A. (2012). The effects of
substituting glassware for plasticware and the use of an ethanol vector on oocyte
maturation in vitro. Veterinary Medicine International, 2012, 914715.
doi:10.1155/2012/914715
Macaulay, A. D., Gilbert, I., Caballero, J., Barreto, R., Fournier, E., Tossou, P., Sirard, M-A,
Clarke, H. J., Khandjian, E. W., Richard, F. J., Hyttel, O., & Robert, C. (2014). The
gametic synapse; RNA transfer to the bovine oocyte. Biology of Reproduction.
doi:10.1095/biolreprod.114.119867
Macaulay, A. D., Hamilton, C. K., King, W. A., & Bartlewski, P. M. (2013). Influence of
physiological concentrations of androgens on the developmental competence and sex ratio
of in vitro produced bovine embryos. Reproductive Biology, 13(1), 41–50.
doi:10.1016/j.repbio.2013.01.175
Machtinger, R., Combelles, C. M. H., Missmer, S. A., Correia, K. F., Williams, P., Hauser, R., &
Racowsky, C. (2013). Bisphenol-A and human oocyte maturation in vitro. Human
Reproduction, 28(10), 2735–45. doi:10.1093/humrep/det312
MacLusky, N. J., Hajszan, T., & Leranth, C. (2005). The Environmental Estrogen Bisphenol A
Inhibits Estradiol-Induced Hippocampal Synaptogenesis. Environmental Health
Perspectives, 113(6), 675–679. doi:10.1289/ehp.7633
Mailhes J.B., Handel M. A., 2008, Chapter 54: Animal Models for Investigating the Causes and
Mechanisms of Mammalian Germ Cell Aneuploidy, In: Sourcebook of Models for
Biomedical Research, Ed. Humana Press, ISSN 978-1-58829-933-8 (Print) 978-1-59745285-4 (Online).
Maître, B., Jornot, L., & Junod, A. F. (1993). Effects of inhibition of catalase and superoxide
dismutase activity on antioxidant enzyme mRNA levels. The American Journal of
Physiology, 265(6 Pt 1), L636–L643.
Majno, G., & Joris, I. (1995). Apoptosis, oncosis, and necrosis. An overview of cell death. The
American Journal of Pathology, 146, 3–15.
Mamo, S., Carter, F., Lonergan, P., Leal, C. L., Al Naib, A., McGettigan, P., Mehta, J. P., Evans,
A. C. O., & Fair, T. (2011). Sequential analysis of global gene expression profiles in
immature and in vitro matured bovine oocytes: potential molecular markers of oocyte
maturation. BMC Genomics, 12(1), 151. doi:10.1186/1471-2164-12-151
155
Manejwala, F. M., Logan, C. Y., & Schultz, R. M. (1991). Regulation of hsp70 mRNA levels
during oocyte maturation and zygotic gene activation in the mouse. Developmental
Biology, 144(2), 301–308. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/2010034
Mangelsdorf, D. J., Thummel, C., Beato, M., Herrlich, P., Schutz, .G, Umesono, K., Blumberg,
B., Kastner, P., Mark, M., Chambon, P. & Evans, R. M. (1995). The nuclear receptor
superfamily: the second decade. Cell, 83, 835–839.
Marei, W. F., Wathes, D. C., & Fouladi-Nashta, A. A. (2010). Impact of linoleic acid on bovine
oocyte maturation and embryo development. Reproduction, 139, 979–988.
doi:10.1530/REP-09-0503
Markey, C. M., Coombs, M. a., Sonnenschein, C., & Soto, A. M. (2003). Mammalian
development in a changing environment: Exposure to endocrine disruptors reveals the
developmental plasticity of steroid-hormone target organs. Evolution and Development, 5,
67–75. doi:10.1046/j.1525-142X.2003.03011.x
Markey, C. M., Wadia, P. R., Rubin, B. S., Sonnenschein, C., & Soto, A. M. (2005). Long-term
effects of fetal exposure to low doses of the xenoestrogen bisphenol-A in the female mouse
genital tract. Biology of Reproduction, 72(6), 1344–1351.
doi:10.1095/biolreprod.104.036301
Marumoto, T., Saya, H., & Zhang, D. (2005). Aurora-A - A guardian of poles. Nature Reviews
Cancer. 5.1, 42.
Matthews, J.B., Twomey, K., & Zacharewski, T.R. (2001). In vitro and in vivo interactions of
bisphenol A and its metabolite, bisphenol A glucuronide, with estrogen receptors α and β.
Chemical Research in Toxicology, 14, 149–157.
Matwee, C., Betts, D. H., & King, W. A. (2000). Apoptosis in the early bovine embryo. Zygote,
8(1), 57–68. doi:10.1017/S0967199400000836
Matwee, C., Kamaruddin, M., Betts, D. H., Basrur, P. K., & King, W. A. (2001). The effects of
antibodies to heat shock protein 70 in fertilization and embryo development. Molecular
Human Reproduction, 7(9), 829–37. Retrieved from
http://www.ncbi.nlm.nih.gov/pubmed/11517289
Matzuk, M. M., Burns, K. H., Viveiros, M. M., & Eppig, J. J. (2002). Intercellular
communication in the mammalian ovary: oocytes carry the conversation. Science, 296,
2178–2180. doi:10.1126/science.1071965
Meeker, J. D., Ferguson, K. K. (2011). Relationship between urinary phthalate and bisphenol A
concentrations and serum thyroid measures in U.S. adults and adolescents from the
National Health and Nutrition Examination Survey (NHANES) 2007-2008. Environmental
Health Perspectives, 119, 1396–1402.
Memili, E., & First, N. L. (1999). Control of gene expression at the onset of bovine embryonic
development, Biology of Reproduction, 61, 1198–1207.
156
Mendez, R., Hake, L. E., Andresson, T., Littlepage, L. E., Ruderman, J. V, & Richter, J. D.
(2000). Phosphorylation of CPE binding factor by Eg2 regulates translation of c-mos
mRNA. Nature, 404(6775), 302–307. doi:10.1038/35005126
Mendiola, J., Jørgensen, N., Andersson, A. M., Calafat, A. M., Ye, X., Redmon, J. B., Drobnis,
E. Z., Wang, C., Sparks, A., Thurston, S. W., Liu, F., & Swan, S. H. (2010). Are
environmental levels of bisphenol A associated with reproductive function in fertile men?
Environmental Health Perspectives, 118, 1286–1291. doi:10.1289/ehp.1002037.
Ménézo, Y. J. R., Chouteau, J., Torelló, M. J., Girard, A., & Veiga, A. (1999). Birth weight and
sex ratio after transfer at the blastocyst stage in humans. Fertility and Sterility, 72(2), 221–
224. doi:10.1016/S0015-0282(99)00256-3
Merchant Research & Consulting (2014). Bisphenol A (BPA): 2015 world market outlook and
forecast up to 2019. Available on: http://mcgroup.co.uk/researches/bisphenol-a-bpa
Metwally, M., Cutting, R., Tipton, a, Skull, J., Ledger, W. L., & Li, T. C. (2007). Effect of
increased body mass index on oocyte and embryo quality in IVF patients. Reproductive
Biomedicine Online, 15(5), 532–538. doi:10.1016/S1472-6483(10)60385-9
Minamiyama, Y., Ichikawa, H., Takemura, S., Kusunoki, H., Naito, Y., & Yoshikawa, T. (2010).
Generation of reactive oxygen species in sperms of rats as an earlier marker for evaluating
the toxicity of endocrine-disrupting chemicals. Free Radical Research, 44(12), 1398–1406.
Mocarelli, P., Gerthoux, P. M., Ferrari, E., Patterson, D. G., Kieszak, S. M., Brambilla, P.,
Vincoli, N., Signorini, S., Tramacere, P., Carreri, V., Sampson, E. J., Turner, W. E., &
Needham, L. L. (2000). Paternal concentrations of dioxin and sex ratio of offspring.
Lancet, 355, 1858–1863. doi:10.1016/S0140-6736(00)02290-X
Modina, S., Borromeo, V., Luciano, a M., Lodde, V., Franciosi, F., & Secchi, C. (2007).
Relationship between growth hormone concentrations in bovine oocytes and follicular
fluid and oocyte developmental competence. European Journal of Histochemistry : EJH,
51, 173–80. Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/17921112
Mok-Lin, E., Ehrlich, S., Williams, P. L., Petrozza, J., Wright, D. L., Calafat, A. M., Ye, X., &
Hauser, R. (2010). Urinary bisphenol A concentrations and ovarian response among
women undergoing IVF. International Journal of Andrology, 33(2), 385–93.
doi:10.1111/j.1365-2605.2009.01014.x
Mørck, T. J., Sorda, G., Bechi, N., Rasmussen, B. S., Nielsen, J. B., Ietta, F., Rytting, E.,
Mathiesen, L., Paulesu, L., & Knudsen, L. E. (2010). Placental transport and in vitro
effects of Bisphenol A. Reproductive Toxicology, 30(1), 131–7.
doi:10.1016/j.reprotox.2010.02.007
Moriya, A. & Hiroshige T. (1978). Sex ratio of offspring of rats bred at 50C. The International
Journal of Biometerology, 22, 312–315.
157
Moriyama, K., Tagami, T., Akamizu, T., Usui, T., Saijo, M., Kanamoto, N., Hataya, Y.,
Shimatu, A., Kuzuya, H. & Nakao, K. (2002). Thyroid hormone action is disrupted by
bisphenol A as an antagonist. The Journal of Clinical Endocrinology and Metabolism,
87(11), 5185–90. doi:10.1210/jc.2002-020209
Mtango, N. R., Potireddy, S., & Latham, K. E. (2008). Oocyte quality and maternal control of
development. International Review of Cell and Molecular Biology, 268(08), 223–90.
doi:10.1016/S1937-6448(08)00807-1
Neuer, A, Mele, C., Liu, H. C., Rosenwaks, Z., & Witkin, S. S. (1998). Monoclonal antibodies to
mammalian heat shock proteins impair mouse embryo development in vitro. Human
Reproduction (Oxford, England), 13(4), 987–990.
Neuer, A., Spandorfer, S. D., Giraldo, P., Jeremias, J., Dieterle, S., Korneeva, I., Liu, H-C.,
Rosenwaks, Z., & Witkin, S. S. (1999). Heat shock protein expression during
gametogenesis and embryogenesis. Infectious Diseases in Obstetrics and Gynecology,
7(C), 10–16. doi:10.1155/S1064744999000034
Nicol, C. J., Harrison, M. L., Laposa, R. R., Gimelshtein, I.L., Wells, P. G. (1995). A teratologic
suppressor role for p53 in benzo[a]pyrene-treated transgenic p53-deficient mice. Nature
Genetics, 10, 181–187.
Niemann, H., & Wrenzycki, C. (2000). Alterations of expression of developmentally important
genes in preimplantation bovine embryos by in vitro culture conditions: Implications for
subsequent development. Theriogenology, 53(99), 21–34. doi:10.1016/S0093691X(99)00237-X
Nishikawa, M., Iwano, H., Yanagisawa, R., Koike, N., Inoue, H., & Yokota, H. (2010). Placental
transfer of conjugated bisphenol A and subsequent reactivation in the rat fetus.
Environmental Health Perspectives, 118(9), 1196–203. doi:10.1289/ehp.0901575
Norimura, T., Nomoto, S., Katsuki, M., Gondo, Y., & Kondo, S. (1996). p53-dependent
apoptosis suppresses radiation-induced teratogenesis. Nature Medecine, 2:577-580.
Norris, R. P., Ratzan, W. J., Freudzon, M., Mehlmann, L. M., Krall, J., Movsesian, M. A., Wang,
H., Ke, H., Nikolaev, V. O., & Jaffe, L. A. (2009). Cyclic GMP from the surrounding
somatic cells regulates cyclic AMP and meiosis in the mouse oocyte. Development, 136,
1869–1878. doi:10.1242/dev.035238
Okada, A. & Kai, O. (2008). Effects of estradiol-17beta and bisphenol A administered
chronically to mice throughout pregnancy and lactation on the male pups’ reproductive
system. Asian Journal of Andrology, 10, 271–276.
Ornoy, A. (2007). Embryonic oxidative stress as a mechanism of teratogenesis with special
emphasis on diabetic embryopathy. Reproductive Toxicology, 24, 31–41.
doi:10.1016/j.reprotox.2007.04.004
158
Ortiz, D. (2009). Developmental changes in zebrafish embryos exposed to bisphenol A leeched
from commonly used plastics. Honors Thesis. Liberty University.
Orvos, H., Kozinszky, Z., & Bartfai, G. (2001). Natural variation in the human sex ratio. Human
Reproduction, 16, 803.
Paczkowski, M., Yuan, Y., Fleming-Waddell, J., Bidwell, C.A., Spurlock, D., & Krisher, R.L.
(2011). Alterations in the transcriptome of porcine oocytes derived from prepubertal and
cyclic females is associated with developmental potential. Journal of Animal Science, 89,
3561–3571.
Palanza, P., Gioiosa, L., vom Saal, F.S., & Parmigiani, S. (2008). Effects of developmental
exposure to bisphenol A on brain and behavior in mice. Environmental Research, 108, 150
157.
Pantaleon, M., Ryan, J. P., Gil, M., & Kaye, P. L. (2001). An unusual subcellular localization of
GLUT1 and link with metabolism in oocytes and preimplantation mouse embryos. Biology
of Reproduction, 64, 1247–1254. doi:10.1095/biolreprod64.4.1247
Park, K., & Kwak, I. S. (2013). Expression of stress response HSP70 gene in Asian paddle crabs,
Charybdis japonica, exposure to endocrine disrupting chemicals, bisphenol A (BPA) and 4nonylphenol (NP). Ocean Science Journal, 48, 207–214. doi:10.1007/s12601-013-0017-y
Pascreau, G., Eckerdt, F., Lewellyn, A. L., Prigent, C., & Maller, J. L. (2009). Phosphorylation
of p53 is regulated by TPX2-Aurora A in Xenopus oocytes. Journal of Biological
Chemistry, 284(9), 5497–5505. doi:10.1074/jbc.M805959200
Pastva, S.D., Villalobos, S.A., Kannan, K., & Giesy, J.P. (2001). Morphological effects of
Bisphenol-A on the early life stages of medaka (Oryzias latipes). Chemosphere, 45, 535
541.
Paula-Lopes, F. F., & Hansen, P. J. (2002a). Apoptosis is an adaptive response in bovine
preimplantation embryos that facilitates survival after heat shock. Biochemical and
Biophysical Research Communications, 295, 37–42. doi:10.1016/S0006-291X(02)00619-8
Paula-Lopes, F. F., & Hansen, P. J. (2002b). Heat shock-induced apoptosis in preimplantation
bovine embryos is a developmentally regulated phenomenon. Biology of Reproduction, 66,
1169–1177. doi:10.1095/biolreprod66.5.1570
Pearce, E. N., & Braverman, L. E. (2009). Environmental pollutants and the thyroid. Best
Practice & Research. Clinical Endocrinology & Metabolism, 23(6), 801–13.
doi:10.1016/j.beem.2009.06.003
Pelech, S., Jelinkova, L., Susor, A., Zhang, H., Shi, X., Pavlok, A., Kubelka, M., & Kovarova, H.
(2008). Antibody Microarray Analyses of Signal Transduction Protein Expression and
Phosphorylation during Porcine Oocyte Maturation research articles, 2860–2871.
159
Peretz, J., Craig, Z. R., & Flaws, J. A. (2012). Bisphenol A Inhibits Follicle Growth and Induces
Atresia in Cultured Mouse Antral Follicles Independently of the Genomic Estrogenic
Pathway. Biology of Reproduction, 87(June), 63–63. doi:10.1095/biolreprod.112.101899
Peretz, J., Neese, S. L., & Flaws, J. A. (2013). Mouse strain does not influence the overall effects
of bisphenol a-induced toxicity in adult antral follicles. Biology of Reproduction, 89(5),
108. doi:10.1095/biolreprod.113.111864
Peretz, J., Vrooman, L., Ricke, W. A., Hunt, P. A., Ehrlich, S., Hauser, R., Padmanabhan, V.,
Taylor, H. S., Swan, S. H., VandeVoort, C. A., & Flaws, J. A. (2014). Bisphenol A and
reproductive health: update of experimental and human evidence, 2007-2013.
Environmental health perspectives, 122(8), 775-786.
Pérez-Crespo, M., Ramírez, M. A., Fernández-González, R., Rizos, D., Lonergan, P., Pintado,
B., & Gutiérrez-Adán, A. (2005). Differential sensitivity of male and female mouse
embryos to oxidative induced heat-stress is mediated by glucose-6-phosphate
dehydrogenase gene expression. Molecular Reproduction and Development, 72(October
2004), 502–510. doi:10.1002/mrd.20366
Pfeiffer, E., Rosenberg, B., Deuschel, S., & Metzler, M. (1997). Interference with microtubules
and induction of micronuclei in vitro by various bisphenols. Mutation Research - Genetic
Toxicology and Environmental Mutagenesis, 390(November 1996), 21–31.
doi:10.1016/S0165-1218(96)00161-9
Pines, J., & Hunter, T. (1989). Isolation of a human cyclin cDNA: evidence for cyclin mRNA
and protein regulation in the cell cycle and for interaction with p34cdc2. Cell, 58(5), 833–
846. doi:10.1016/0092-8674(89)90936-7
Piper, E. K., Jackson, L. A, Bagnall, N. H., Kongsuwan, K. K., Lew, A. E., & Jonsson, N. N.
(2008). Gene expression in the skin of Bos taurus and Bos indicus cattle infested with the
cattle tick, Rhipicephalus (Boophilus) microplus. Veterinary Immunology and
Immunopathology, 126(1-2), 110–9. doi:10.1016/j.vetimm.2008.06.011
Planelló, R., Martínez-Guitarte, J. L., & Morcillo, G. (2008). The endocrine disruptor bisphenol
A increases the expression of HSP70 and ecdysone receptor genes in the aquatic larvae of
Chironomus riparius. Chemosphere, 71, 1870–1876.
doi:10.1016/j.chemosphere.2008.01.033
Pocar, P., Augustin, R., Gandolfi, F., & Fischer, B. (2003). Toxic effects of in vitro exposure to
p-tert-octylphenol on bovine oocyte maturation and developmental competence. Biology of
Reproduction, 69(2), 462–8. doi:10.1095/biolreprod.102.010355
Pocar, P., Brevini, T. A., Perazzoli, F., Cillo, F., Modina, S., & Gandolfi, F. (2001). Cellular and
molecular mechanisms mediating the effects of polychlorinated biphenyls on oocyte
developmental competence in cattle. Molecular Reproduction and Development,
60(September 2000), 535–541. doi:10.1002/mrd.1118
160
Pratt, N. C., & Lisk, R. D. (1989). Effects of social stress during early pregnancy on litter size
and sex ratio in the golden hamster (Mesocricetus auratus). Journal of Reproduction and
Fertility, 87, 763–769.
Pritchett, J. J., Kuester, R. K., & Sipes, I. G. (2002). Metabolism of bisphenol a in primary
cultured hepatocytes from mice, rats, and humans. Drug Metabolism and Disposition: The
Biological Fate of Chemicals, 30(11), 1180–1185.
Purcell, S. H., & Moley, K. H. (2009). Glucose transporters in gametes and preimplantation
embryos. Trends in Endocrinology and Metabolism, 20(10), 483–489.
doi:10.1016/j.tem.2009.06.006
Qin, F., Wang, L., Wang, X., Liu, S., Xu, P., Wang, H., Wu, T., Zhang, Y., Zheng, Y., Li, M.,
Zhang, X., Yuan, C., Hu, G., & Wang, Z. (2013). Bisphenol A affects gene expression of
gonadotropin-releasing hormones and type I GnRH receptors in brains of adult rare
minnow Gobiocypris rarus. Comparative Biochemistry and Physiology. Toxicology &
Pharmacology : CBP, 157(2), 192–202. doi:10.1016/j.cbpc.2012.11.002
Rama Raju, G.A., Prakash, G.J., Krishna, K.M., & Madan, K. (2007). Meiotic spindle and zona
pellucida characteristics as predictors of embryonic development: a preliminary study
using PolScope imaging. Reproductive Biomedicine Online, 14, 166–174.
Ramakrishnan, S. & Wayne, N. L. (2008). Impact of bisphenol-A on early embryonic
development and reproductive maturation. Reproductive Toxicology (Elmsford, N.Y.),
25(2), 177–83. doi:10.1016/j.reprotox.2007.11.002
Ramos, J. G., Varayoud, J., Kass, L., Rodríguez, H., Costabel, L., Muñoz-De-Toro, M., &
Luque, E. H. (2003). Bisphenol A induces both transient and permanent histofunctional
alterations of the hypothalamic-pituitary-gonadal axis in prenatally exposed male rats.
Endocrinology, 144(February), 3206–3215. doi:10.1210/en.2002-0198
Rastinejad, F., Huang, P., Chandra, V., & Khorasanizadeh, S. (2013). Understanding nuclear
receptor form and function using structural biology. Journal of Molecular Endocrinology,
51(3), T1–T21. doi:10.1530/JME-13-0173
Rho, G., Balasubramanian, S., Kim, D., Son, W., Cho, S., Kim, J., Kumar, M., & Choe, S.
(2007). Influence of In Vitro Oxygen Concentrations on Preimplantation Embryo
Development , Gene Expression and Production of Hanwoo Calves Following Embryo
Transfer, 496(January 2006), 486–496. doi:10.1002/mrd
Richter JD. (1996). Dynamics of poly(A) addition and removal during development. In: Hershey
J, Mathews M, Sonenberg N, editors. Translational control. New York: Cold Spring
Harbor Laboratory Press, 481–503.
Richter, C. A, Birnbaum, L. S., Farabollini, F., Newbold, R. R., Rubin, B. S., Talsness, C. E.,
Vandenberg, J. G., Walser-Kuntz, D. R., vom Saal, F. S. (2007). In vivo effects of
bisphenol A in laboratory rodent studies. Reproductive Toxicology, 24(2), 199–224.
doi:10.1016/j.reprotox.2007.06.004
161
Rizos, D., Ward, F., Duffy, P., Boland, M. P., & Lonergan, P. (2002). Consequences of bovine
oocyte maturation, fertilization or early embryo development in vitro versus in vivo:
Implications for blastocyst yield and blastocyst quality. Molecular Reproduction and
Development, 61, 234–248. doi:10.1002/mrd.1153
Rjasanowski, I., Klöting, I., & Kovacs, P. (1998). Decreased prevalence of atopy in rheumatoid
arthritis Altered sex ratio in offspring of mothers with insulin-dependent diabetes mellitus
Nosocomial transmission of dengue from a needlestick injury Treatment of cutaneous
leishmaniasis in Colombia with daps, 351, 497–498.
Robker, R. L., Akison, L. K., Bennett, B. D., Thrupp, P. N., Chura, L. R., Russell, D. L., Lane,
M., & Norman, R. J. (2009). Obese women exhibit differences in ovarian metabolites,
hormones, and gene expression compared with moderate-weight women. Journal of
Clinical Endocrinology and Metabolism, 94, 1533–1540. doi:10.1210/jc.2008-2648
Rodriguez-Martinez, H. (2012). Assisted reproductive techniques for cattle breeding in
developing countries: A critical appraisal of their value and limitations. Reproduction in
Domestic Animals, 47, 21–26. doi:10.1111/j.1439-0531.2011.01961.x
Rubin, B. S. (2011). Bisphenol A: an endocrine disruptor with widespread exposure and multiple
effects. The Journal of Steroid Biochemistry and Molecular Biology, 127(1-2), 27–34.
doi:10.1016/j.jsbmb.2011.05.002
Rubin, B. S., Murray, M. K., Damassa, D. A., King, J. C., & Soto, A. M. (2001). Perinatal
exposure to low doses of bisphenol A affects body weight, patterns of estrous cyclicity,
and plasma LH levels. Environmental Health Perspectives, 109(7), 675–680.
doi:10.1289/ehp.01109675
Russell, D.F., Baqir, S., Bordignon, J. & Betts, D.H. (2006). The impact of oocyte maturation
media on early bovine embryonic development. Molecular Reproduction and
Development, 73, 1255–1270.
Sagirkaya, H., Misirlioglu, M., Kaya, A., First, N. L., Parrish, J. J., & Memili, E. (2007).
Developmental potential of bovine oocytes cultured in different maturation and culture
conditions. Animal Reproduction Science, 101(3-4), 225–240.
doi:10.1016/j.anireprosci.2006.09.016
Saito, K., Furukawa, E., Kobayashi, M., Fukui, E., Yoshizawa, M., & Matsumoto, H. (2014).
Degradation of estrogen receptor α in activated blastocysts is associated with implantation
in the delayed implantation mouse model. Molecular Human Reproduction, 20(5), 384–91.
doi:10.1093/molehr/gau004
Sakurai, K., Kawazuma, M., Adachi, T., Harigaya, T., Saito, Y., Hashimoto, N., & Mori, C.
(2004). Bisphenol A affects glucose transport in mouse 3T3-F442A adipocytes. British
Journal of Pharmacology, 141(2), 209–14. doi:10.1038/sj.bjp.0705520
162
Salian, S., Doshi, T., & Vanage, G. (2009). Neonatal exposure of male rats to bisphenol A
impairs fertility and expression of Sertoli cell junctional proteins in the testis. Toxicology,
265, 56–67.
Sanfins, A., Lee, G. Y., Plancha, C. E., Overstrom, E. W., & Albertini, D. F. (2003). Distinctions
in meiotic spindle structure and assembly during in vitro and in vivo maturation of mouse
oocytes. Biology of Reproduction, 69(6), 2059–67. doi:10.1095/biolreprod.103.020537
Sar, M., & Welsch, F. (1999). Differential expression of estrogen receptor-beta and estrogen
receptor-alpha in the rat ovary. Endocrinology, 140(2), 963–971.
doi:10.1210/endo.140.2.6533
Sartori, R., Sartor-Bergfelt, R., Mertens, S. A., Guenther, J. N., Parrish, J. J., & Wiltbank, M. C.
(2002). Fertilization and early embryonic development in heifers and lactating cows in
summer and lactating and dry cows in winter. Journal of Dairy Science, 85(11), 2803–
2812. doi:S0022-0302(02)74367-1 [pii]\n10.3168/jds.S0022-0302(02)74367-1
Scheil, V., Zu, A., & Ko, H. (2009). Embryo Development, Stress Protein (Hsp70) Responses ,
and Histopathology in Zebrafish (Danio rerio) Following Exposure to Nickel Chloride ,
Chlorpyrifos, and Binary Mixtures of Them, 83–93. doi:10.1002/tox
Schier, A. F. (2007). The maternal-zygotic transition: death and birth of RNAs. Science (New
York, N.Y.), 316(5823), 406–7. doi:10.1126/science.1140693
Schirling, M., Jungmann, D., Ladewig, V., Ludwichowski, K. U., Nagel, R., Köhler, H. R., &
Triebskorn, R. (2006). Bisphenol A in artificial indoor streams: II. Stress response and
gonad histology in Gammarus fossarum (Amphipoda). Ecotoxicology, 15, 143–156.
doi:10.1007/s10646-005-0044-2
Schönfelder, G., Wittfoht, W., Hopp, H., Talsness, C.E., Paul, M., & Chahoud, I. (2002). Parent
bisphenol A accumulation in the human maternal-fetal-placental unit. Environmental
Health Perspectives, 110, A703–A707.
Schultz, R. M. (2005). From egg to embryo: A peripatetic journey. Reproduction, 130, 825–828.
doi:10.1530/rep.1.00902
Schultz, R.M. (1993). Regulation of zygotic gene activation in the mouse. Bioessays, 15, 531–
538.
Shelby, M. D. (2008). NTP-CERHR monograph on the potential human reproductive and
developmental effects of bisphenol A. NTP CERHR MON, 22: v, vii–ix, 1–64.
Shuda, K., Schindler, K., Ma, J., Schultz, R. M., & Donovan, P. J. (2009). Aurora kinase B
modulates chromosome alignment in mouse oocytes. Molecular Reproduction and
Development, 76(11), 1094–1105. doi:10.1002/mrd.21075
Silva, C. F., Sartorelli, E. S., Castilho, A. C. S., Satrapa, R. a., Puelker, R. Z., Razza, E. M., …
Barros, C. M. (2013). Effects of heat stress on development, quality and survival of Bos
163
indicus and Bos taurus embryos produced in vitro. Theriogenology, 79, 351–357.
doi:10.1016/j.theriogenology.2012.10.003
Sirard, M. A., Richard, F., Blondin, P., & Robert, C. (2006). Contribution of the oocyte to
embryo quality. Theriogenology, 65, 126–136. doi:10.1016/j.theriogenology.2005.09.020
Spindler, R. E., Pukazhenthi, B. S., & Wildt, D. E. (2000). Oocyte metabolism predicts the
development of cat embryos to blastocyst in vitro. Molecular Reproduction and
Development, 56(August 1999), 163–171. doi:10.1002/(SICI)10982795(200006)56:2<163::AID-MRD7>3.0.CO;2-3
Stebbins-Boaz, B., Hake, L. E., & Richter, J. D. (1996). CPEB controls the cytoplasmic
polyadenylation of cyclin, Cdk2 and c-mos mRNAs and is necessary for oocyte maturation
in Xenopus. The EMBO Journal, 15(10), 2582–2592. doi:8665866
Stewart, Z.A. & Pietenpol, J.A. (2001). P53 signalling and cell cycle checkpoints. Chem Res
Toxicol. 14, 243-263.
Su, Y. Q., Sugiura, K., Woo, Y., Wigglesworth, K., Kamdar, S., Affourtit, J., & Eppig, J. J.
(2007). Selective degradation of transcripts during meiotic maturation of mouse oocytes.
Developmental Biology, 302(1), 104–117. doi:10.1016/j.ydbio.2006.09.008
Suchanek, E., Simunic, V., Macas, E., Kopjar, B., & Grizelj, V. (1988): Prostaglandin F2 alpha,
progesterone and estradiol concentrations in human follicular fluid and their relation to
success of in vitro fertilization. The European Journal of Obstetrics & Gynecology and
Reproductive Biology, 28, 331-339
Sugiura, K., & Eppig, J. J. (2005). Society for Reproductive Biology Founders’ Lecture 2005
Control of metabolic cooperativity between oocytes and their companion granulosa cells
by mouse oocytes. Reproduction, Fertility and Development, 17, 667–674.
doi:10.1071/RD05071
Susiarjo, M., Hassold, T. J., Freeman, E., & Hunt, P. A. (2007). Bisphenol A exposure in utero
disrupts early oogenesis in the mouse. PLoS Genetics, 3(1), e5.
doi:10.1371/journal.pgen.0030005
Susiarjo, M., Sasson, I., Mesaros, C., & Bartolomei, M. S. (2013). Bisphenol A exposure
disrupts genomic imprinting in the mouse. PLoS Genetics, 9(4), e1003401.
doi:10.1371/journal.pgen.1003401
Suzuki, A., Sugihara, A., Uchida, K., Sato, T., Ohta, Y., Katsu, Y., Watanabe, H., & Iguchi, T.
(2002). Developmental effects of perinatal exposure to bisphenol-A and diethylstilbestrol
on reproductive organs in female mice. Reproductive Toxicology, 16, 107–116.
Swain, J. E. (2010). Optimizing the culture environment in the IVF laboratory: Impact of pH and
buffer capacity on gamete and embryo quality. Reproductive BioMedicine Online, 21(1),
6–16. doi:10.1016/j.rbmo.2010.03.012
164
Swain, J. E., Ding, J., Wu, J., & Smith, G. D. (2008). Regulation of spindle and chromatin
dynamics during early and late stages of oocyte maturation by aurora kinases. Molecular
Human Reproduction, 14(5), 291–299. doi:10.1093/molehr/gan015
Tabuchi, Y., Zhao, Q.-L., & Kondo, T. (2002). DNA microarray analysis of differentially
expressed genes responsive to bisphenol A, an alkylphenol derivative, in an in vitro mouse
Sertoli cell model. Japanese Journal of Pharmacology, 89, 413–416. doi:10.1254/jjp.89.413
Takahashi, M. (2012). Oxidative stress and redox regulation on in vitro development of
mammalian embryos. The Journal of Reproduction and Development, 58(1), 1–9.
Retrieved from http://www.ncbi.nlm.nih.gov/pubmed/22450278
Takahashi, M., Mukai, H., Oishi, K., Isagawa, T., & Ono, Y. (2000). Association of immature
hypophosphorylated protein kinase C; an Anchoring protein CG-NAP. Journal of
Biological Chemistry, 275, 34592–34596. doi:10.1074/jbc.M005285200
Takahashi, T., Takahashi, E., Igarashi, H., Tezuka, N., & Kurachi, H. (2003). Impact of
oxidative stress in aged mouse oocytes on calcium oscillations at fertilization. Molecular
Reproduction and Development, 66, 143–152.
Takao, T., Nanamiya, W, Nazarloo, H. P., Matsumoto, R., Asaba, K., Hashimoto, K. (2003).
Exposure to the environmental estrogen bisphenol A differentially modulated estrogen
receptor-alpha and -beta immunoreactivity and mRNA in male mouse testis. Life Sciences,
72(10), 1159-1169.
Takai, Y., Tsutsumi, O., Ikezuki, Y., Kamei, Y., Osuga, Y., Yano, T., & Taketan, Y. (2001).
Preimplantation exposure to bisphenol A advances postnatal development. Reproductive
Toxicology, 15, 71–74. doi:10.1016/S0890-6238(00)00119-2
Takamiya, K., Kostourou, V., Adams, S., Jadeja, S., Chalepakis, G., Scambler, P. J., Huganir, R.
L. & Adams, R. H. (2004). A direct functional link between the multi-PDZ domain protein
GRIP1 and the Fraser syndrome protein Fras1. Nature Genetics, 36(2), 172–177.
doi:10.1038/ng1292
Takeuchi, T., Tsutsumi, O., Ikezuki, Y., Takai, Y., & Taketani, Y. (2004). Positive relationship
between androgen and the endocrine disruptor, bisphenol A, in normal women and women
with ovarian dysfunction. Endocrine Journal, 51(2), 165–9. Retrieved from
http://www.ncbi.nlm.nih.gov/pubmed/15118266
Tamura, H., Takasaki, A., Miwa, I., Taniguchi, K., Maekawa, R., Asada, H., Taketani, T.,
Matsuoka, A., Yamagata, Y., Shimamura, K., Morioka, H., Ishikawa, H., Reiter, R. J. &
Sugino, N. (2008). Oxidative stress impairs oocyte quality and melatonin protects oocytes
from free radical damage and improves fertilization rate. Journal of Pineal Research, 44,
280–287. doi:10.1111/j.1600-079X.2007.00524.x
Tanaka, Y., Kanai, Y., Okada, Y., Nonaka, S., Takeda, S., Harada, A., & Hirokawa, N. (1998).
Targeted disruption of mouse conventional kinesin heavy chain, kif5B, results in abnormal
165
perinuclear clustering of mitochondria. Cell, 93(7), 1147–1158. doi:10.1016/S00928674(00)81459-2
Tarin, J. (1996). Potential effects of age-associated oxidative stress on mammalian
oocytes/embryos. Molecular Human Reproduction, 2, 717–724.
Tatemoto, H., Sakurai, N., & Muto, N. (2000). Protection of porcine oocytes against apoptotic
cell death caused by oxidative stress during In vitro maturation: role of cumulus cells.
Biology of Reproduction, 63(3), 805–810. doi:10.1095/biolreprod63.3.805
Tatone, C., Amicarelli, F., Carbone, M. C., Monteleone, P., Caserta, D., Marci, R., Artini, P. G.,
Piomboni, P., & Focarelli, R. (2008). Cellular and molecular aspects of ovarian follicle
ageing. Human Reproduction Update, 14(2), 131–142. doi:10.1093/humupd/dmm048
Taylor, D. M., Handyside, A H., Ray, P. F., Dibb, N. J., Winston, R. M., & Ao, A. (2001).
Quantitative measurement of transcript levels throughout human preimplantation
development: analysis of hypoxanthine phosphoribosyl transferase. Molecular Human
Reproduction, 7(2), 147–154.
Telford, N. A., Watson, A. J., & Schultz, G. A. (1990). Transition from maternal to embryonic
control in early mammalian development: a comparison of several species. Molecular
Reproduction and Development, 26, 90-100.
Thompson JG, Simpson AC, PughCW,Donnelley PE, Tervit HR. 1990. Effect of oxygen
concentration on in vitro development of preim- plantation sheep and cattle embryos.
Journal of Reproduction and Fertility, 89, 573–578.
Tiffin, G.J., Rieger, D., Betteridge, K.J., Yadav, B.R. & King, W.A. (1991). Glucose and
glutamine metabolism in pre-attachment cattle embryos in relation to sex and stage of
development. Journal of Reproduction and Fertility, 93, 125–132.
Tiwari, D. & Vanage, G. (2013). Mutagenic effect of bisphenol A on adult rat male germ cells
and their fertility. Reproductive Toxicology, 40, 60–68.
doi:10.1016/j.reprotox.2013.05.013.
Tomari, H., Honjou, K., Nagata, Y., & Horiuchi, T. (2011). Relationship between meiotic
spindle characteristics in human oocytes and the timing of the first zygotic cleavage after
intracytoplasmic sperm injection. Journal of Assisted Reproduction and Genetics, 28(11),
1099–104. doi:10.1007/s10815-011-9634-5
Trdan Lušin, T., Roškar, R., & Mrhar, A. (2012). Evaluation of bisphenol A glucuronidation
according to UGT1A1*28 polymorphism by a new LC-MS/MS assay. Toxicology, 292(1),
33–41. doi:10.1016/j.tox.2011.11.015
Trinei, M., Giorgio, M., Cicalese, A., Barozzi, S., Ventura, A., Migliaccio, E., Milia, E., Padura,
I. M., Raker, V. A., Maccarana M., Petronilli, V., Minucci, S., Bernardi, P., Lanfrancone,
L., & Pelicci, P. G. (2002). A p53-p66Shc signalling pathway controls intracellular redox
166
status, levels of oxidation-damaged DNA and oxidative stress-induced apoptosis.
Oncogene, 21(24), 3872–3878. doi:10.1038/sj.onc.1205513
Trivers, R.L. & Willard, D.E. (1973). Natural selection of parental ability to vary the sex ratio of
offspring. Science, 179, 90–92.
Trounson, A., Anderiesz, C. & Jones, G. (2001). Maturation of human oocytes in vitro and their
developmental competence. Reproduction,121, 51–75.
Tsutsui, T., Tamura, Y., Yagi, E., Hasegawa, K., Takahashi, M, Maizumi, N, Yamaguchi, F., &
Barrett, J. C. (1998). Bisphenol-A induces cellular transformation, aneuploidy and DNA
adduct formation in cultured Syrian hamster embryo cells. The International Journal of
Cancer, 75(2), 290–294.
Uzbekova, S., Arlot-Bonnemains, Y., Dupont, J., Dalbiès-Tran, R., Papillier, P., Pennetier, S.,
Thelie, A., Perreau, C., Mermillod, P., Prigent, C., & Uzbekov, R. (2008). Spatio-temporal
expression patterns of aurora kinases a, B, and C and cytoplasmic polyadenylationelement-binding protein in bovine oocytes during meiotic maturation. Biology of
Reproduction, 78(2), 218–233. doi:10.1095/biolreprod.107.061036
Valckx, S. D. M., Arias-Alvarez, M., De Pauw, I., Fievez, V., Vlaeminck, B., Fransen, E., Pols,
P. E. J., & Leroy, J. L. M. R. (2014). Fatty acid composition of the follicular fluid of
normal weight, overweight and obese women undergoing assisted reproductive treatment:
a descriptive cross-sectional study. Reproductive Biology and Endocrinology : RB&E,
12(1), 13. doi:10.1186/1477-7827-12-13
Valentino, R., D’Esposito, V., Passaretti, F., Liotti, A., Cabaro, S., Longo, M., Perruolo, G.,
Oriente, F., Beguinot, F. & Formisano, P. (2013). Bisphenol-A impairs insulin action and
up-regulates inflammatory pathways in human subcutaneous adipocytes and 3T3-L1 cells.
PLoS ONE, 8(12), 1–10. doi:10.1371/journal.pone.0082099
Van Blerkom, J., Davis, P. W., & Lee, J. (1995). ATP content of human oocytes and
developmental potential and outcome after in-vitro fertilization and embryo transfer.
Human Reproduction (Oxford, England), 10(2), 415–424.
van de Leemput, E.E., Vos, PLAM, Zeinstra, E.C., Bevers, M.M., der Weijden, G.C., Dieleman,
S.J. (1999). Improved in vitro embryo development using in vivo matured oocytes from
heifers superiovulated with a controlled preovulatory LH surge. Theriogenology, 52, 335
349.
Van der Auwera, I., & D’Hooghe, T. (2001). Superovulation of female mice delays embryonic
and fetal development. Human Reproduction (Oxford, England), 16(6), 1237–1243.
Van Hoeck, V., Leroy, J. L. M. R., Alvarez, M. A., Rizos, D., Gutierrez-Adan, a., Schnorbusch,
K., Bols, P. E. J., Leese, H. J. & Sturmey, R. G. (2013). Oocyte developmental failure in
response to elevated nonesterified fatty acid concentrations: Mechanistic insights.
Reproduction, 145, 33–44. doi:10.1530/REP-12-0174
167
Vandaele, L., Thys, M., Bijttebier, J., Van Langendonckt, a., Donnay, I., Maes, D., Meyer, E., &
Van Soom, A. (2010). Short-term exposure to hydrogen peroxide during oocyte maturation
improves bovine embryo development. Reproduction, 139(3), 505–511. doi:10.1530/REP09-0430
Vandenberg, L. N., Ehrlich, S., Belcher, S. M., Ben-Jonathan, N., Dolinoy, D. C., Hugo, E. R.,
Hunt, P. A., Newbold, R. R., Rubin, B. S., Saili, K. S., Soto, A. M., Wang, H-S., vom Saal,
F. S. (2013). Low dose effects of bisphenol A: An integrated review of in vitro, laboratory
animal, and epidemiology studies. Endocrine Disruptors, 1(February 2015), 0–1.
doi:10.4161/endo.26490
Vandenberg, L. N., Hauser, R., Marcus, M., Olea, N., & Welshons, W. V. (2007). Human
exposure to bisphenol A (BPA). Reproductive Toxicology, 24(2), 139–77.
doi:10.1016/j.reprotox.2007.07.010
Vandenberg, L. N., Maffini, M. V, Sonnenschein, C., Rubin, B. S., & Soto, A. M. (2009).
Bisphenol-A and the great divide: a review of controversies in the field of endocrine
disruption. Endocrine Reviews, 30(1), 75–95. doi:10.1210/er.2008-0021
Varghese, A. C., Ly, K. D., Corbin, C., Mendiola, J., & Agarwal, A. (2010). Oocyte
developmental competence and embryo development: impact of lifestyle and
environmental risk factors. Reproductive Biomedicine Online, 22(5), 410–420.
doi:10.1016/j.rbmo.2010.11.009
Vasquez, Y. M., & DeMayo, F. J. (2013). Role of nuclear receptors in blastocyst implantation.
Seminars in Cell and Developmental Biology, 24(0), 724–735.
doi:10.1016/j.semcdb.2013.08.004
Vigneault, C., Gravel, C., Vallée, M., McGraw, S., & Sirard, M.-A. (2009). Unveiling the bovine
embryo transcriptome during the maternal-to-embryonic transition. Reproduction
(Cambridge, England), 137(2), 245–57. doi:10.1530/REP-08-0079
Virant-klun, I., Knez, K., Tomazevic, T., & Skutella, T. (2013). Gene Expression Profiling of
Human Oocytes Developed and Matured In Vivo or In Vitro, 2013.
Volarcik, K., Sheean, L., Goldfarb, J., Woods, L., Abdul-Karim, F. W., & Hunt, P. (1998). The
meiotic competence of in-vitro matured human oocytes is influenced by donor age:
Evidence that folliculogenesis is compromised in the reproductively aged ovary. Human
Reproduction, 13(1), 154–160. doi:10.1093/humrep/13.1.154
Voronina, E., & Wessel, G. M. (2003). The regulation of oocyte maturation. Current Topics in
Developmental Biology, 58(401), 53–110. Retrieved from
http://www.ncbi.nlm.nih.gov/pubmed/14711013
Wang, W. H., & Keefe, D. L. (2002). Prediction of chromosome misalignment among in vitro
matured human oocytes by spindle imaging with the PolScope. Fertility and Sterility,
78(5), 1077–1081. doi:10.1016/S0015-0282(02)04196-1
168
Wang, W., Craig, Z. R., Basavarajappa, M. S., Hafner, K. S., & Flaws, J. a. (2012). Mono (2ethylhexyl) Phthalate Induces Oxidative Stress and Inhibits Growth of Mouse Ovarian
Antral Follicles. Biology of Reproduction, 87(October), 1–10.
doi:10.1095/biolreprod.112.102467
Wang, W., Meng, L., Hackett, R. J. … (2002). Rigorous thermal control during intracytoplasmic
sperm injection stabilizes the meiotic spindle and improves fertilization and pregnancy
rates, 77(6), 1274–1277.
Watson, A. J., Sousa, P. De, Caveney, A., Barcroft, L. C., Natale, D., Urquhart, J., & Westhusin,
M. E. (2000). Impact of Bovine Oocyte Maturation Media on Oocyte Transcript Levels,
Blastocyst Development, Cell Number , and Apoptosis 1, 364, 355–364.
Watson, a. J., Westhusin, M. E., De Sousa, P. a., Betts, D. H., & Barcroft, L. C. (1999). Gene
expression regulating blastocyst formation. Theriogenology, 51(1), 117–133.
doi:10.1016/S0093-691X(98)00236-2
Wei, X., Huang, Y., Wong, M. H., Giesy, J. P., & Wong, C. K. C. (2011). Assessment of risk to
humans of bisphenol A in marine and freshwater fish from Pearl River Delta, China.
Chemosphere, 85(1), 122–8. doi:10.1016/j.chemosphere.2011.05.038
Weisskopf, M. G., Anderson, H. A., & Hanrahan, L. P. (2003). Decreased sex ratio following
maternal exposure to polychlorinated biphenyls from contaminated Great Lakes sportcaught fish: a retrospective cohort study. Environmental Health : A Global Access Science
Source, 2, 2. doi:10.1186/1476-069X-2-2
Welch, W. J. (1992). Mammalian stress response: cell physiology, structure/function of stress
proteins, and implications for medicine and disease. Physiological Reviews, 72(4), 1063–
1081.
Welch, W. J., & Feramisco, J. R. (1984). Nuclear and nucleolar localization of the 72,000-dalton
heat shock protein in heat shocked mammalian cells. Journal of Biological Chemistry,
259(7), 4501–4513.
Wells, D., & Patrizio, P. (2008). Gene expression profiling of human oocytes at different
maturational stages and after in vitro maturation. American Journal of Obstetrics and
Gynecology, 198(4). doi:10.1016/j.ajog.2007.12.030
Wells, D., Alfarawati, S., & Fragouli, E. (2008). Use of comprehensive chromosomal screening
for embryo assessment: microarrays and CGH. Molecular Human Reproduction, 14(12),
703–10. doi:10.1093/molehr/gan062
Welshons, W. V, Nagel, S. C., & vom Saal, F. S. (2006). Large effects from small exposures. III.
Endocrine mechanisms mediating effects of bisphenol A at levels of human exposure.
Endocrinology, 147(6 Suppl), S56–69. doi:10.1210/en.2005-1159
Welshons, W. V., Thayer, K. a., Judy, B. M., Taylor, J. A., Curran, E. M., & vom Saal, F. S.
(2003). Large effects from small exposures. I. Mechanisms for endocrine-disrupting
169
chemicals with estrogenic activity. Environmental Health Perspectives, 111(8), 994–1006.
doi:10.1289/ehp.5494
Wessel, G. M., Brooks, J. M., Green, E., Haley, S., Voronina, E., Wong, J., Zaydfudim, V., and
Conner, S. (2001). The biology of cortical granules. International Review of Cytology,
209, 117–206.
Wetherill, Y. B., Akingbemi, B. T., Kanno, J., McLachlan, J. A, Nadal, A., Sonnenschein, C.,
Watson, C. S., Zoeller, R. T., & Belcher, S. M. (2007). In vitro molecular mechanisms of
bisphenol A action. Reproductive Toxicology (Elmsford, N.Y.), 24(2), 178–98.
doi:10.1016/j.reprotox.2007.05.010
Wilson, G. N., Howe, M., & Stover, J. M. (1985). Delayed developmental sequences in rodent
diabetic embryopathy. Pediatric Research, 19(12), 1337–1340.
Wolstenholme, J. T., Edwards, M., Shetty, S. R. J., Gatewood, J. D., Taylor, J. a, Rissman, E. F.,
& Connelly, J. J. (2012). Gestational exposure to bisphenol a produces transgenerational
changes in behaviors and gene expression. Endocrinology, 153(8), 3828–38.
doi:10.1210/en.2012-1195
Wolstenholme, J. T., Rissman, E. F., & Connelly, J. J. (2011). The role of Bisphenol A in
shaping the brain, epigenome and behavior. Hormones and Behavior, 59(3), 296–305.
doi:10.1016/j.yhbeh.2010.10.001
Wood, T. C., & Wildt, D. E. (1997). Effect of the quality of the cumulus-oocyte complex in the
domestic cat on the ability of oocytes to mature, fertilize and develop into blastocysts in
vitro. Journal of Reproduction and Fertility, 110(2), 355–360. doi:10.1530/jrf.0.1100355
Wrenzycki, C., Herrmann, D., & Niemann, H. (2007). Messenger RNA in oocytes and embryos
in relation to embryo viability. Theriogenology, 68(SUPPL. 1), 77–83.
doi:10.1016/j.theriogenology.2007.04.028
Wrenzycki, C., Herrmann, D., Carnwath, J. W., & Niemann, H. (1999). Alterations in the
relative abundance of gene transcripts in preimplantation bovine embryos cultured in
medium supplemented with either serum or PVA. Molecular Reproduction and
Development, 53(November 1998), 8–18. doi:10.1002/(SICI)10982795(199905)53:1<8::AID-MRD2>3.0.CO;2-K
Wrenzycki, C., Herrmann, D., Carwath, J.W., Niemann, H. (1998). Expression of RNA from
developmentally important genes in preimplantation bovine embryos produced in TCM
supplemented with BSA. Journal of Reproduction and Fertility, 112, 387–398.
Wrenzycki, C., Herrmann, D., Keskintepe, L., Martins, A., Sirisathien, S., Brackett, B., &
Niemann, H. (2001). Effects of culture system and protein supplementation on mRNA
expression in pre-implantation bovine embryos. Human Reproduction (Oxford, England),
16(5), 893–901.
170
Xiao, S., Diao, H., Smith, M. A., Song, X., & Ye, X. (2011). Preimplantation exposure to
bisphenol A (BPA) affects embryo transport, preimplantation embryo development, and
uterine receptivity in mice. Reproductive Toxicology, 32(4), 434–441.
doi:10.1016/j.reprotox.2011.08.010
Yamasita M, Mita K, Yoshida N, Kondo T. (2000). Molecular mechanisms of the initiation of
oocyte maturation: General and species-specific aspects. Progress in Cell Cycle Research,
4, 115–129
Yan, C., Wang, P., DeMayo, J., DeMayo, F. J., Elvin, J. A., Carino, C., Prasad, S. V., Skinner, S.
S. Dunbar, B. S., Dube, J. L., Celeste, A. J. & Matzuk, M. M. (2001). Synergistic roles of
bone morphogenetic protein 15 and growth differentiation factor 9 in ovarian function.
Molecular Endocrinology (Baltimore, Md.), 15(March), 854–866.
doi:10.1210/mend.15.6.0662
Yan, S., Song, W., Chen, Y., Hong, K., Rubinstein, J., Wang, H.S. (2013). Low-dose bisphenol
A and estrogen increase ventricular arrhythmias following ischemia- reperfusion in female
rat hearts. Food and Chemical Toxicology.
Yang, F., Lei, X., Rodriguez-palacios, A., Tang, C., & Yue, H. (2013). Selection of reference
genes for quantitative real-time PCR analysis in chicken embryo fibroblasts infected with
avian leukosis virus subgroup J. BMC Research Notes, 6(1), 1. doi:10.1186/1756-0500-6402
Yang, L., Lv, J., Wang, X., Zhang, J., Li, Q., Zhang, T., Zhang, Z., & Zhang, L. (2015). Direct
interactions in the recognition between the environmental estrogen bisphenol AF and
human serum albumin. Journal of Molecular Recognition. doi: 10.1002/jmr.2463.
Yao, L.-J., Zhong, Z.-S., Zhang, L.-S., Chen, D.-Y., Schatten, H., & Sun, Q.-Y. (2004). AuroraA is a critical regulator of microtubule assembly and nuclear activity in mouse oocytes,
fertilized eggs, and early embryos. Biology of Reproduction, 70(5), 1392–1399.
doi:10.1095/biolreprod.103.025155
Ye, J., Coleman, J., Hunter, M. G., Craigon, J., Campbell, K. H. S., & Luck, M. R. (2007).
Physiological temperature variants and culture media modify meiotic progression and
developmental potential of pig oocytes in vitro. Reproduction (Cambridge, England),
133(5), 877–86. doi:10.1530/REP-06-0318
Ye, X., Pierik, F.H., Angerer, J., Meltzer, H.M., Jaddoe, V.W., Tiemeier, H., Hoppin, J. A., &
Longnecker, M. P. (2009). Levels of metabolites of organophosphate pesticides,
phthalates, and bisphenol A in pooled urine specimens from pregnant women participating
in the Norwegian Mother and Child Cohort Study (MoBa). The International Journal of
Hygiene and Environmental Health, 212, 481 491.
Ying, C., Hsu, W. L., Hong, W. F., Cheng, W. T. K., & Yang, Y. C. (2000). Estrogen receptor is
expressed in pig embryos during preimplantation development. Molecular Reproduction
171
and Development, 55(May 1999), 83–88. doi:10.1002/(SICI)10982795(200001)55:1<83::AID-MRD11>3.0.CO;2-V
Yoshihara, S., Mizutare, T., Makishima, M., Suzuki, N., Fujimoto, N., Igarashi, K., & Ohta, S.
(2004). Potent estrogenic metabolites of bisphenol A and bisphenol B formed by rat liver
S9 fraction: Their structures and estrogenic potency. Toxicological Sciences, 78, 50–59.
doi:10.1093/toxsci/kfh047
Yuan, Y., Ida, J. M., Paczkowski, M., & Krisher, R. L. (2011). Identification of developmental
competence-related genes in mature porcine oocytes. Molecular Reproduction and
Development, 78(8), 565–575. doi:10.1002/mrd.21351
Zalko, D., Soto, A.M., Dolo, L., Dorio, C., Rathahao, E., Debrauwer, L., Faure, R., Cravedi, J.P.,
2003. Biotransformations of bisphenol A in a mammalian model: answers and new
questions raised by low-dose metabolic fate studies in pregnant CD1 mice. Environmental
Health Perspectives 111, 309–319.
Zhang, Q., Li, Q., Li, J., Li, X., Liu, Z., Song, D., & Xie, Z. (2008). b-DAZL: A novel gene in
bovine spermatogenesis. Progress in Natural Science, 18(10), 1209–1218.
doi:10.1016/j.pnsc.2008.02.012
Zhang, X. F., Zhang, L. J., Feng, Y. N., Chen, B., Feng, Y. M., Liang, G. J., Li, L., & Shen, W.
(2012). Bisphenol A exposure modifies DNA methylation of imprint genes in mouse fetal
germ cells. Molecular Biology Reports, 39(9), 8621–8628. doi:10.1007/s11033-012-17167
Zheng, P., Si, W., Bavister, B. D., Yang, J., Ding, C., & Ji, W. (2003). 17β-Estradiol and
progesterone improve in-vitro cytoplasmic maturation of oocytes from unstimulated
prepubertal and adult rhesus monkeys. Human Reproduction, 18(10), 2137–2144.
doi:10.1093/humrep/deg410
Zoeller, R. T., Bansal, R., & Parris, C. (2005). Bisphenol-A, an environmental contaminant that
acts as a thyroid hormone receptor antagonist in vitro, increases serum thyroxine, and alters
RC3/neurogranin expression in the developing rat brain. Endocrinology, 146(2), 607–12.
doi:10.1210/en.2004-1018
172
`