Embed Size (px)
Citation preview
MCL-1 - GUARDIAN OF THE OVARIAN RESERVE: CHARACTERIZATION OF THE CONTRIBUTION OF
MCL-1 TO OOCYTE SURVIVAL
by
SHAKIB OMARI
A THESIS SUBMITTED IN CONFORMITY WITH THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY
DEPARTMENT OF PHYSIOLOGY UNIVERSITY OF TORONTO
© Copyright by Shakib Omari 2014
ii
MCL-1 – GUARDIAN OF THE OVARIAN RESERVE:
CHARACTERIZATION OF THE CONTRIBUTION OF MCL-1 TO
OOCYTE SURVIVAL
Shakib Omari
Doctor of Philosophy
Department of Physiology
University of Toronto
2014
ABSTRACT
Oocyte quality and the maintenance of the ovarian reserve are essential factors in reproductive
competence and fertility. The predominant portion of ovarian follicles are lost to natural death
and the disruption of factors involved in conservation of the oocyte pool result in an untimely
follicle exhaustion known as premature ovarian failure. Additionally, advancing maternal age is
accompanied by accelerated follicle loss and the production of oocytes with poor developmental
competence. Thus, identification of factors regulating oocyte quality and survival are essential in
management and preservation of fertility.
The anti-apoptotic Bcl-2 family member MCL-1 plays a pro-survival role in various cell types;
however, its contribution to oocyte survival is unknown. In this thesis we characterized the
phenotype caused by oocyte-specific Mcl-1-deficiency, and determined its role in the
maintenance of the primordial follicle pool, growing oocyte survival and oocyte quality.
Disruption of Mcl-1 resulted in a premature exhaustion of the ovarian reserve, characterized by
early primordial follicle loss. The increasingly diminished surviving cohort of growing oocytes
possessed elevated markers of autophagy and mitochondrial dysfunction. Ovulated Mcl-1
deficient oocytes displayed an increased susceptibility to cellular fragmentation with elevated
iii
activation of the apoptotic cascade. Concomitant deletion of the pro-apoptotic Bax rescued the
primordial follicle phenotype and ovulated oocyte death, but did not impact mitochondrial
dysfunction associated with Mcl-1-deficiency.
Furthermore, we assessed the impact of the cytokine Kit ligand and metabolic supplementation,
both associated with increased oocyte survival, on MCL-1 expression. We identified MCL-1 as a
pro-survival target of these pathways in oocytes. Diminished Mcl-1-levels, both induced as well
as physiologically driven, coincide with impaired mitochondrial output; resulting in starvation-
induced autophagy and poor ovarian reserve. We classify MCL-1 as the essential survival factor
required for maintenance of the primordial follicle pool, growing follicle survival and effective
oocyte mitochondrial bioenergetics.
iv
ACKNOWLEDGMENTS
This thesis is dedicated to my parents, who are always there for me in every moment of need,
and without whom none of this would have been possible.
I would like to thank my supervisor, Dr. Andrea Jurisicova for all her support and guidance over
the years, and to my committee members Dr. Robert Casper, Dr. Norman Rosenblum and Dr.
Helen McNeill for their help and direction.
I would also like to extend my thanks to Dr. Ted Brown and Dr. Ian Rogers, for their
encouragement and their positivity and their consistent willingness to help out whenever I
reached out to them.
On a personal and professional level I would like to thank all the following people who have
helped me out at all stages of my PhD, and without whom these last years would have been dull
and lifeless. Firstly, I want to thank Dr. Alexandra Kollara for always being there whenever I
needed any sort of help, whether it was listening to my presentations, or just lending a helpful ear
in a moment of need. I would also like to thank Drs Shadab Bhai, Premy and Shawn for their
friendship over the years. We’ve been through a lot. All my fellow labmates also need
mentioning, for all the late nights at work ordering pizza, to the random runs to nearest bar when
it just got too crazy. Sreca- you’re amazing, Tuchka- you’re too great, squash partner Krusty,
Russanthy and Legs- you’re awesome, Talon- stop stressing, Richard, Aluet, Han, Scrubs, Nella,
Mangow, MJ, Peep, Gibby, Abhi, Jacqui, Fatima, I couldn’t have done any of this without you
all!
Finally, I’d like to thank my family, my parents and my sis, for their love and support. Thanks
Shaz, for everything! BL… 143!! And a huge heartfelt thanks to all the friends I made in Toronto
that were always there for me whenever my studies had its ups and downs!
v
CONTRIBUTIONS
The following people have contributed to the data presented in various portions of this thesis
Dr. Joseph Opferman , St. Jude Children’s Research Hospital, kindly provided us with the floxed
Mcl-1 mice.
Dr. Jonathan Tilly, and Dr. Razq Hakem kindly supplied the Bax and Bim deficient mice (used in
Chapter 2).
Metabolite levels for ATP, Citrate, Fumarate and Malate (referred to in Chapter 2 and 3) were
generously analyzed by the lab of Dr. Kelle Moley
Histomorphometric analyses (in Chapter 2) of PN7, PN14, PN21, and PN90 ovaries (follicle
counts) were performed by Sarah Cao and especially Mara Waters
Work presented in this thesis was supported by CIHR
vi
TABLE OF CONTENTS
Table of Contents
ACKNOWLEDGMENTS ................................................................................................................ iv
CONTRIBUTIONS ......................................................................................................................... v
TABLE OF CONTENTS ................................................................................................................. vi
LIST OF FIGURES .......................................................................................................................... x
LIST OF APPENDICES ................................................................................................................ xii
ABBREVIATIONS ....................................................................................................................... xiii
1 LITERATURE REVIEW ........................................................................................................... 1
1.1 OVERVIEW ....................................................................................................................... 1
1.2 OVARIAN DEVELOPMENT ............................................................................................ 2
1.2.1 Oogenesis and Folliculogenesis .............................................................................. 2
1.2.2 Initial Recruitment .................................................................................................. 5
1.2.3 Cyclic Recruitment ............................................................................................... 10
1.2.4 Initial and Cyclic Recruitment Factors Associated With Premature Ovarian
Failure ................................................................................................................... 12
1.3 OOCYTE-GRANULOSA INTERCOMMUNICATION ................................................. 13
1.3.1 Extra-Cellular Signaling ....................................................................................... 13
1.3.2 Gap Junction Mediated Signaling ......................................................................... 13
1.4 OOCYTE METABOLISM ............................................................................................... 15
1.5 MITOCHONDRIA ........................................................................................................... 18
1.5.1 Mitochondrial Structure and Metabolism ............................................................. 18
1.5.2 Oocyte Mitochondria ............................................................................................ 19
1.5.3 Mitochondria and Oocyte Developmental Competence ....................................... 20
1.6 PROGRAMMED CELL DEATH TYPE 1 ...................................................................... 21
vii
1.6.1 Intrinsic Pathway of Apoptosis ............................................................................. 22
1.7 PROGRAMMED CELL DEATH TYPE 1 IN THE OVARY ......................................... 26
1.7.1 Pro-apoptotic Bcl-2 Family in the Ovary .............................................................. 26
1.7.2 Anti-apoptotic Bcl-2 Family in the Ovary ............................................................ 29
1.8 PROGRAMMED CELL DEATH TYPE 2 ...................................................................... 31
1.8.1 Autophagosome Formation ................................................................................... 34
1.8.2 Formation of Autolysosome and Substrate Degradation ...................................... 35
1.9 AUTOPHAGY (PCD TYPE 2) IN THE OVARY ........................................................... 36
1.9.1 Bcl-2 Family in Autophagy and Metabolism ........................................................ 37
1.10 MCL-1 ............................................................................................................................... 38
1.10.1 Gene, Transcript and Protein Structure ................................................................. 39
1.10.2 Transcriptional Regulation of Mcl-1 .................................................................... 39
1.10.3 Post-Translational Regulation of MCL-1 ............................................................. 40
1.10.4 Metabolic Role of MCL-1 .................................................................................... 43
1.10.5 Autophagic Role of MCL-1 .................................................................................. 44
1.10.6 MCL-1 in the Ovary .............................................................................................. 45
1.11 THESIS HYPOTHESIS AND OBJECTIVES ................................................................. 46
2 ASSESSING THE ROLE OF ANTI-APOPTOTIC BCL-2 MEMBER MCL-1 IN
OOCYTE AND FOLLICLE FATE ......................................................................................... 48
2.1 INTRODUCTION ............................................................................................................ 48
2.2 MATERIALS AND METHODS ...................................................................................... 51
2.2.1 Animals ................................................................................................................. 51
2.2.2 Collection of MII and GV Oocytes ....................................................................... 52
2.2.3 Histological Analyses ........................................................................................... 53
2.2.4 TUNEL Assays ..................................................................................................... 55
2.2.5 Ovulation Rates, Fragmentation Rates and Breeding Performance ...................... 55
viii
2.2.6 Mitochondrial Analyses – Live cell stains ............................................................ 56
2.2.7 Immunofluorescence Staining .............................................................................. 57
2.2.8 Imaging ................................................................................................................. 59
2.2.9 Metabolic Profile .................................................................................................. 59
2.2.10 Statistics ................................................................................................................ 59
2.3 RESULTS ......................................................................................................................... 60
2.3.1 Breeding Performance, Ovulation Rates and Histomorphometric Analyses ........ 61
2.3.2 Markers of Apoptosis in Growing Follicle Pool ................................................... 68
2.3.3 Markers of Autophagy in Growing Follicle Pool ................................................. 69
2.3.4 Mitochondrial Functionality in Ovulated Oocyte Pool ......................................... 73
2.3.5 Viability of Ovulated Oocytes .............................................................................. 80
2.3.6 Rescue of Mcl-1-Deficient Phenotype by Deletion of Bax .................................. 81
2.4 DISCUSSION ................................................................................................................... 88
3 CYTOKINE AND METABOLIC REGULATION OF MCL-1 FUNCTION IN MURINE
OOCYTES ............................................................................................................................... 93
3.1 INTRODUCTION ............................................................................................................ 93
3.2 MATERIALS AND METHODS ...................................................................................... 96
3.2.1 Animals ................................................................................................................. 96
3.2.2 Collection of GV Oocytes ..................................................................................... 97
3.2.3 Collection of Growing Oocyte Pool – PI3 Kinase Pathway ................................. 97
3.2.4 Collection of Growing Oocyte Pool – Pyruvate Treatment .................................. 98
3.2.5 Treatment with Inhibitors of Pyruvate Uptake and Fatty Acid Breakdown ......... 98
3.2.6 Co-Immunoprecipitations ..................................................................................... 99
3.2.7 Western Blots, Antibodies, Reagents .................................................................... 99
3.2.8 Measurement of ATP and Lipid Droplets ........................................................... 100
3.2.9 Statistics .............................................................................................................. 100
ix
3.3 RESULTS ....................................................................................................................... 100
3.3.1 Impact of KL-Activated PI3 Kinase Pathway Stimulation on MCL-1 ............... 101
3.3.2 Prevention of Oocyte Death with Pyruvate Supplementation ............................ 102
3.3.3 Importance of Mcl-1 in Oocyte Metabolism. ..................................................... 104
3.3.3 Regulation of Energy Output in Mcl-1-deficient oocytes ................................... 109
3.3.4 Alternative Means of Energy Production ........................................................... 112
3.4 DISCUSSION ................................................................................................................. 115
4 OVERALL DISCUSSION ..................................................................................................... 121
4.1 Regulation of Primordial Follicle Fate ........................................................................... 123
4.2 Primordial Follicle Growth and Growing Follicle Survival ........................................... 127
4.3 Additional Mitochondrial Role for MCL-1 (MCL-1Matrix
) ............................................. 131
4.4 Meiotic Resumption and Ovulation ................................................................................ 136
5 REFERENCES ....................................................................................................................... 141
APPENDIX ................................................................................................................................. 163
x
LIST OF FIGURES
Figure 1.1 Initial and Cyclic Recruitment during Normal Follicular Development. ............. 4
Figure 1.2 Molecular Pathways involved in Initial and Cyclic Recruitment. ......................... 7
Figure 1.3. Intrisic Pathway of Apoptosis. ................................................................................ 25
Figure 1.4. Molecular Pathways of Autophagy. ....................................................................... 33
Figure 1.5. Mcl-1 mRNA Transcript and Protein Structure. ................................................ 42
Figure 2.1. Patterns of MCL-1 Expression and Verification of Mcl-1 Oocyte-Specific
Excision. ............................................................................................................................... 63
Figure 2.2. Breeding Performance, Ovulation Rates and Histomorphometric Analyses of
Mcl-1cKO Females and Controls. ..................................................................................... 66
Figure 2.3. Histomorphometric Analyses of Mcl-1cKO Females and Controls. ................... 67
Figure 2.4. Markers of Apoptosis and Autophagy in GV Oocytes. ........................................ 72
Figure 2.5. Markers of Autolysosome Formation in GV Oocytes. ......................................... 75
Figure 2.6. Markers of Mitochondrial Functionality. ............................................................. 77
Figure 2.7. Markers of Mitochondrial Functionality, DNA Damage and Spindle Assembly.
............................................................................................................................................... 79
Figure 2.8. Markers of Autophagy and Apoptosis in MII Oocytes. ....................................... 83
Figure 2.9. Rescue of Mcl-1-Deficient Follicle Loss by Concurrent Bax-Ablation. .............. 86
Figure 2.10. Impact of Bax-Ablation on Mcl-1-Deficient Oocyte Mitochondrial Function
and Apoptosis. ..................................................................................................................... 87
xi
Figure 3.1. Activation/Inhibition of PI3 Kinase Pathway and Impact on MCL-1 Expression.
............................................................................................................................................. 103
Figure 3.2. Impact of Pyruvate Treatment on Oocyte Survival and MCL-1 Expression. . 106
Figure 3.3. Impact of Starvation on Oocyte Survival in Mcl-1cKO. .................................... 108
Figure 3.4. Co-Immunoprecipitation with MCL-1 Pulldown in Ovarian Lysates. ............ 111
Figure 3.5. Lipid Droplet Formation in Mcl-1cKO and Controls. ....................................... 113
Figure 4.1. Overview of Mechanisms Involved in Oocyte Survival or Death via Regulation
of MCL-1, Presented in this Thesis. ................................................................................ 140
xii
LIST OF APPENDICES
Figure A1. Histomorphometric Analyses of BimKO and Protection Against Radiation-
Induced Primordial Follicle Death. ................................................................................. 163
Figure A2. Assessing Impact of γ-Irradiation on MCL-1 and BIM Expression and MCL-1-
BIM Interaction. ............................................................................................................... 165
xiii
ABBREVIATIONS
4EBP eukaryotic initiation factor 4E binding proteins
aa amino acid
Acetyl coA acetyl coenzyme A
ADP adenosine diphosphate
ANOVA analysis of variance
AMP adenosine monophosphate
AMPK adenosine-monophosphate protein activated-kinase
AP alkaline phosphatase
ATG autophagy-related
ATP adenosine triphosphate
ATPAF ATP-synthase assembly factor
Bcl-2 B-cell lymphoma 2
Bcl2l- bcl-2-like-
BH bcl-2 homology
BMP15 bone-morphogenetic protein 15
BOD bcl-2-related ovarian death
BOK bcl-2-related ovarian killer
BSA bovine serum albumin
cAMP cyclic adenosine monophosphate
Caspase cysteinyl aspartic acid proteases
cGMP cyclic guanosine monophosphate
C-KIT kit receptor
cKO conditional knockout
COC cumulus-oocyte-complex
Co-IP co-immunoprecipitation
CPT-1 carnitine palmitoyl-transferase-1
CX connexin
DAB diamino-benzidine
DAPI 4',6-diamidino-2-phenylindole
DDX4 dead box polypeptide 4
DISC death-inducing signaling complex
DMEM dulbeccos modified eagles medium
DMSO dimethyl sulfoxide
DNA deoxyribonucleic acid
dpc days post coitum
xiv
EDTA ethylenediaminetetraacetic acid
EGF epidermal growth factor
ERK extracellular signal-regulated kinases
FAD/FADH2 flavin adenine dinucleotide
FADD fas-associated death domain
FAS fatty acid synthetase ligand
Fig. Figure
FITC fluorescein isothiocyanate
FOXO forkhead transcription factor subclass O
FSH follicle stimulating hormone
g centrifugal force unit
GDF-9 growth differentiation factor 9
GSK-3 glycogen synthase kinase 3
GV germinal vesicle
GWAS genome wide association studies
HCA hydoxy-cinnamic acid
hCG human chorionic gonadotropin
hr hour
HTF human tubal fluid
IBMX 3-isobutyl-1-methylxanthine
IGF-1 insulin-like growth factor-1
IHC immuno-histochemistry
IMM inner mitochondrial membrane
IMS inter-membrane space
IP immuno-precipitation
IVM in vitro maturation
JNK cJun N-terminal kinase
KL kit ligand
kb kilobase
KO knockout
LH luteinizing hormone
LC3 microtubule-associated protein 1 light chain
LIR lc-3-interacting region
LAMP1/2 lysosome-associated membrane proteins 1 and 2
xv
mg milligram
ug microgram
ml milliliter
ul microliter
um micrometer
mm millimeter
mM millimolar
uM micromolar
MCL-1 myeloid cell leukemia 1
MCL-1S myeloid cell leukemia 1-short
MCL-1ES myeloid cell leukemia 1-extra short
MEF mouse embryonic fibroblasts
mHTF modified human tubal fluid
MII metaphase II
min minute
MPP mitochondrial processing peptidase
mRNA messenger ribonucleic acid
mtDNA mitochondrial deoxyribonucleic acid
mTOR mammalian target of rapamycin
mTORc1 mammalian target of rapamycin complex 1
MULE mcl-1 ubiquitin ligase E3
ng nanogram
nm nanometer
nM nanomolar
NAD/NADH nicotinamide adenine dinucleotide
NAD(P)H nicotinamide adenine dinucleotide phosphate
Neo neomycin cassette
OMM outer mitochondrial membrane
p70S6K p70 ribosomal s6 kinase
PAH polycyclic aromatic hydrocarbon
PBS phosphate buffered saline
PCD programmed cell death
PDE3A phosphodiestrase
PDHA1 pyruvate dehydrogenase alpha 1
PDK1 phopsphoinositide dependent kinase 1
PE phophotidylethanolamine
PEP phosphoenolpyruvate
PEST proline, glutamic acid, serine, threonine
xvi
PI3K phosphotidylinositol 3 kinase (class1 or class3)
PIP2 phosphotidylinositol 3,4,5 diphosphate
PIP3 phosphotidylinositol 3,4,5 triphosphate
PKA protein kinase a
PKB protein kinase b
PMSG pregnant mare serum gonadotropin
PN postnatal
POF premature ovarian failure
PTEN phospatase and tensin homolog deleted on chromosome 10
PVDF polyvinylidene difluoride
rpS6 ribosomal protein s6
RAPTOR regulatory associated protein of mammalian target of rapamycin
ROS reactive oxygen species
RNA ribonucleic acid
rpm revolutions per minute
RIPA radioimmunoprecipitation assay
SCF stem cell factor
Ser serine
SEM standard error of mean
SDS sodium dodecyl sulfate
TCA tricarboxylic acid cycle
TEM transmission electron microscopy
Thr threonine
TM transmembrane
TNF tumor necrosis factor
TOM20 translocase of outer mitochondrial membrane 20kDA
TRADD tumor necrosis factor receptor-associated death domain
TRAIL tumor necrosis factor-related apoptosis-inducing ligand
TRITC tetramethylrhodamine-5(and-6)-isothiocyanate
TSC1/2 tuberous sclerosis 1 and 2
TX triton x
U unit
ULK unc51-like kinase
USP9X ubiquitin-specific peptidase 9 X-linked
WB western blot
ZP3 zona pellucida 3
1
1 LITERATURE REVIEW
1.1 OVERVIEW
The majority of germ cells that are present in the fetal ovary do not survive to ovulation. In fact,
estimates show that 99.9% of germ cells are eliminated via activation of programmed cell death
(PCD) [1, 2]. This extensive follicle loss demarcates a normal ovarian lifespan, but premature
depletion of the ovarian follicle pool can also occur to further exhaust the follicle reserve. These
can include genetic or acquired factors that either sharply diminish or altogether eliminate the
original germ cell supply, or disrupt normal follicular dynamics and growth. Premature Ovarian
Failure (POF) is a syndrome characterized by premature exhaustion of the follicular pool or
disruption of proper follicular development. This condition affects around 1% of all women,
however this number rises to about 30% for women with family history of POF [3].
Maternal age is an additional factor that contributes to the diminished quality of the oocyte and
the resulting embryo [4]. Older oocytes and zygotes have been well documented to have
increased rates of aneuploidies, oxidative damage, mitochondrial and chromosomal
abnormalities and increased fragmentation rates [4-7].
To fully comprehend the mechanics governing normal folliculogenesis, oocyte quality and
factors disrupting them in cases of premature oocyte depletion, it is important to gain an
appreciation for the pathways that regulate early ovarian development, the maintenance of the
dormant ovarian pool, and the recruitment and eventual ovulation of the chosen follicle. In doing
so, it is vital to study factors that control various means of follicular survival, oocyte metabolism,
intercommunication between the oocyte and its surrounding support-cell lineage, and pathways
that govern selection for growth and ovulation. These various aspects can contribute directly to
2
oocyte quality and functionality, and their disruption may result in an increased predilection to
undergo cell death.
Normal ovarian development and the production of a functionally competent oocyte and embryo
is a complex and multifaceted process. Recent studies have alleged the possible maintenance of
ovarian germ stem cells in the adult ovary [8], however the exactitude of these studies have yet
to be fully demonstrated.
1.2 OVARIAN DEVELOPMENT
1.2.1 Oogenesis and Folliculogenesis
Primordial germ cells in the mouse are derived from precursor cells at around 7 days post coitum
(7dpc) (embryonic day 7) and migrate to the undifferentiated gonad, proliferating throughout the
journey [9]. Whereas differentiation of these germ cells into spermatogonia occurs upon
activation of the male pathway, with expression of sex determining region of chromosome Y
(Sry) around 9.5dpc; the absence of the male pathway, in addition to somatic cell factors are
believed to feminize the germ cells in the developing ovary [10]. Incomplete cytokinesis of these
proliferating ‘oogonia’ allows for the maintenance of cytoplasmic bridges between multiple cells
[11, 12], forming cord-like structures akin to germ cell nests, until a collaborative entry into
meiosis around 14.5dpc [13]. Shortly thereafter, the germ cells, now termed ‘oocytes’ after the
onset of meiosis, arrest in the diplotene stage of prophase I. Before birth, the selective
elimination of a subset of oocytes, in addition to somatic cell infiltration of oogonial nests allows
for the individualization of the remaining oocytes with a single layer of flat epithelial granulosa
cells. These individualized units are termed primordial follicles [11-17]. Estimates show that by
3
birth, in a majority of mammalian species, almost two thirds of the germ cell pool has been lost
via atresia of oogonia and oocytes. In fact, as reviewed by Morita et al. [1], during a life time the
pool of ovarian follicles suffers a continuous loss. Those primordial follicles that remain are
maintained in a dormant state until individual follicles are selected from the pool and begin to
grow. The initiation of follicle growth is believed to be due to the synergistic activation of
members of the PI3Kinase and the mTOR pathway [18]. We will explore these pathways in
greater detail ahead.
As reviewed by Edson et al., and demarcated in Figure 1.1 (Fig. 1.1), once the primordial follicle
(classified as type 1) is selected to grow, oocytes of these follicles increase in size and the
surrounding granulosa cells transition from squamous to cuboidal and proliferate, increasing in
number and successive layers around the oocyte [19]. After the initial oocyte growth phase and
transition of the granulosa cells to a single layer of cuboidal cells around the oocyte, the resultant
follicle is termed a primary follicle (type 2). Secondary follicles (type 3), have larger oocytes and
two to four layers of granulosa cells. Around the time of secondary follicle formation, stromal
cells are recruited to the follicle where they differentiate into steroidogenic theca [20]. Pre-antral
(type 4) follicles are characterized by four to six layers of granulosa cells, and with differentiated
theca that become increasingly sensitive to Luteinizing Hormone (LH), developing increasing
numbers of LH receptors on the cell surface. Follicles with five or more layers of granulosa cells
and the presence of an antrum (a pool of follicular fluid) are termed Antral (type 5) follicles.
Prior to pubertal onset, all antral follicles undergo atresia. After puberty, circulating levels of
Follicle Stimulating Hormone (FSH) are able to rescue these antral follicles, and recruit them for
ovulation. Additionally, LH is able to fully activate steroidogenic enzymes in the theca cells
which synthesize androgens that are then transported to the granulosa cells where they are
converted into estradiol, a precursor of estrogen, by the enzyme aromatase.
4
Figure 1.1 Initial and Cyclic Recruitment during Normal Follicular Development.
Primordial germ cells in the fetal ovary (oogonia) are maintained in germ cell ‘nest’ with shared
cytoplasmic bridges. Oogonia death and somatic (granulosa) cell infiltration isolate germ cells into units
termed primordial follicles, composed of a single oocyte surrounded by flat granulosa cells. The neonatal
ovary is stocked with a pool of these primordial follicles, many of which die shortly after birth. The
remaining primordial follicles either remain dormant in the absence of a growth signal, die, or upon
growth factor stimulation (termed Initial Recruitment) begin to grow. Follicle growth is associated with
an increase in oocyte size, in addition to transition of granulosa cells from squamous (flat) to cuboidal and
an increase in successive granulosa cell layers. The transition from the preantral to antral follicle is
characterized by the formation of pools of follicular fluid, believed to be essential for nutrient and oxygen
supply. In the pre-pubertal ovary, all antral follicles undergo atresia. The onset of puberty is accompanied
by the stimulatory effects of Follicle Stimulating Hormone (FSH) that rescues antral follicles and prepares
them for ovulation (termed Cyclic Recruitment).
5
McGee et al. have termed these two instances of follicle recruitment as Initial and Cyclic
recruitment [21]. Initial recruitment refers to the recruitment of primordial follicles from the
dormant pool, and the consequential increase in oocyte size, and layers of cuboidal granulosa. In
rodents, the duration of initial follicular growth can be variable, but the time taken for the
primordial follicle to reach the secondary follicle stage generally exceeds 30days. Furthermore
secondary follicle growth till the antral follicle stage lasts 28days, but prior to puberty, all
instances of initial recruitment result in the death of selected oocytes due to lack of FSH support.
Cyclic recruitment refers to the post-pubertal recruitment of antral follicles from the growing
follicle pool, in preparation for ovulation. This stage is shorter (2-3 days) and an intricate
network of cellular signals regulates the number of follicles progressing through each stage.
Class 1A Phosphotidylinositol 3 Kinases (PI3K) have been found to play a role in both initial
and cyclic recruitment and the molecular mechanisms have been delineated in Figure 1.2 (Fig.
1.2).
1.2.2 Initial Recruitment
The forkhead transcription factor subclass O Foxo3, a member of the PI3 Kinase pathway, when
ablated, was found to cause de-repressed primordial follicle activation [22]; thus implying a role
for Foxo3 in the maintenance of oocyte dormancy. Thereafter, a large number of PI3 Kinase
pathway members were studied to fully characterize the role of this signaling cascade in oocyte
initial recruitment. Zheng et al. have reviewed, in detail, the various members of the PI3 Kinase
signaling cascade and the ovarian phenotype(s) caused by their cell-specific deletion in a variety
of mouse models [23]. In addition to total knockout mouse models, a variety of Cre
Recombinase-mediated conditional mouse knockout models were utilized to assess oocyte-
specific phenotypes. The use of the
6
7
Figure 1.2 Molecular Pathways involved in Initial and Cyclic Recruitment.
Both Initial and Cyclic Recruitment utilize Class 1A PI3 Kinases to regulate survival, proliferation and
growth in oocytes and granulosa cells, respectively. PI3 Kinases are activated by Receptor Protein
Tyrosine Kinases and phosphorylate Phosphotidylinositol 3,4,5 diphosphate (PIP2) to produce PIP3. This
reaction is reversed by the activity of PTEN. PIP3 binds to PDK1 which has been shown to phosphorylate
a number of substrates, two of which include AKT and S6K. AKT Kinase itself has been attributed to the
phosphorylation of a number of downstream targets. These include transcription factors FOXO1 and
FOXO3, Glycogen synthase kinase 3 (GSK-3), and Tsc2. Active FOXO1 and FOXO3 have been
associated with increased cell-cycle arrest and cell death through transcriptional regulation of downstream
factors. GSK-3 is involved in the phosphorylation of a variety of downstream targets regulating
metabolism, proliferation and survival. Phosphorylation of TSC2 causes the inhibition of proper
TSC1/TSC2 complex formation. This inhibition allows for formation of mTORc1 (mTOR complex 1)
which is composed of mTOR, regulatory associated protein of mTOR (Raptor), and a number of other
factors (MLST8, PRAS40, DEPTOR). mTORc1 activity leads to the phosphorylation of a number of
factors, specifically eukaryotic initiation factor 4E binding proteins (4EBPs) and S6K [24]. S6K
phosphorylates rpS6, which together with 4EBP’s, are translational regulators that can increase overall
protein synthesis. Further transcriptional, translational and post-translational regulation of the Bcl-2 anti-
apoptotic factor MCL-1 (orange arrows) has been proposed through various in vitro studies.
8
transgene Tg(Gdf9-Cre)5092-Coo (Gdf-9-Cre) [25], utilizing the promoter region of Growth
differentiation factor 9 (Gdf-9), allows for oocyte-specific excision as early as the primordial
follicle stage, as does the inducible oocyte-specific excision using the transgene Tg(Ddx4-
Cre/ERT2)1Dcas (Vasa-CreERT2
) [26], with an inducible CRE regulated by the DEAD box
polypeptide 4 (Ddx4/Vasa) promoter; whereas the transgene Tg(Zp3-Cre)93Knw (Zp3-Cre) [27]
utilizes the promoter region of Zona Pellucida 3 (ZP3) and results in excision in oocytes of
primary stage follicles and later.
Oocyte-deficient mouse models of Phosphatase and Tensin homolog deleted on chromosome 10
(Pten) driven by Zp3-Cre were phenotypically normal [28], however when crossed with Gdf-9-
Cre or Vasa-CreERT2
, resulted in deregulated primordial follicle activation [26, 29]. The Pten:
Gdf-9-Cre phenotype was rescued by Gdf-9-Cre mediated oocyte-specific deletion of
Phosphoinositide dependent kinase 1 (Pdk1) [30], which by itself caused POF through
accelerated primordial follicle loss. These outcomes are both believed to be due to deregulation
of ribosomal protein S6 (rpS6)-mediated protein translation. Primary oocyte deletion of Pdk1
using Zp3-Cre resulted in normal ovarian development [31], indicating that these two opposing
factors regulate oocyte fate at the primordial follicle stage alone. The total knockout of Protein
Kinase B (PKB/Akt) isoform Akt1 was found to lead to increased oocyte degeneration with
additional granulosa cell defects resulting in POF [32]. Gdf-9-Cre-mediated oocyte-specific
deletion of tuberous sclerosis 1 and 2 (Tsc1, Tsc2) and the total ablation of Foxo3 caused total
primordial follicle activation and the resultant POF [18, 22, 33]. Deletion of Foxo3 using Vasa-
CreERT2
resulted in primordial follicle activation only postnatally [26]. Finally, oocyte-specific
ablation of rpS6 via Gdf-9-Cre resulted in premature primordial follicle and growing follicle loss
[30]. Adhikari et al. demonstrated that the oocyte-specific deletion of Tsc1, resulting in
deregulated primordial follicle activation, was essentially rescued by concurrent deletion of Pdk1
9
[18]. These authors determined that this rescue was due to the ability of both the Mammalian
target of Rapamycin (mTOR) and the PI3 Kinase pathway to phosphorylate p70 Ribosomal S6
Kinase (S6K) at two different residues, leading to rpS6 activation, increased protein translation
and increased primordial follicle activation. Thus, controlled primordial follicle activation and
growth is a collaborative action of mTOR and PI3 Kinase pathway-mediated increases in protein
translation, and the PI3 Kinase-mediated FOXO3-directed release of cell cycle arrest.
A number of growth factors have been postulated to lead to the activation of the PI3 Kinase
pathway and regulate primordial follicle activation and growth. One of the leading candidates
has always been the cytokine Kit ligand (KL) (also referred to as Stem Cell Factor (SCF)),
produced by granulosa cells, which upon binding to its receptor tyrosine kinase C-KIT, has been
shown to activate components of the PI3 Kinase pathway and suppress FOXO3 activity [34]. In
vitro culture of neonatal ovaries with KL, in addition to the use of blocking peptides to prevent
KL binding, have suggested the requirement for KL in primordial follicle activation [35].
Furthermore, genetic studies using various mutations of this ligand and its receptor have resulted
in a number of phenotypes in the ovary, some of which lead to extremely severe oocyte loss and
disrupted folliculogenesis [36, 37]. Ex vivo studies on the isolated follicle pool from post natal
day 8 (PN8) mice have also verified KL activation of the PI3 Kinase pathway, in addition to its
resultant inhibition of Glycogen Synthase Kinase 3 (GSK-3) [38]; a kinase with multiple roles in
proliferation, migration, apoptosis and metabolism [39], yet with limited verified oocyte targets.
Recent evidence, however, has demonstrated that by mutation of the PI3 Kinase binding residue,
KL may not be directly involved in primordial follicle activation, rather it has been relegated to
roles in survival of primordial follicles and the primary to secondary transition [40].
Furthermore, although the impact of KL on follicle survival has been established, the actual
10
mechanistic pathway and factors involved in regulating this survival, downstream of KL, have
yet to be elucidated.
1.2.3 Cyclic Recruitment
In the pre-pubertal ovary, following initial recruitment and progression of the follicle to the
antral stage, the entirety of the antral follicle pool undergoes atresia. It is not until after puberty
that FSH signaling is able to rescue the antral follicles and recruit them for ovulation. This is
termed cyclic recruitment. Deletion of Fsh-β or the Fsh-receptor in granulosa cells has revealed
no change in the ability of the follicle to reach the antral stage, however it does display a
significant impairment in progression beyond the antral stage, upon fertilization and in early
zygote development [41, 42].
As reviewed in Richards et al., FSH signaling is required for activation of a number of pathways
in granulosa cells [43]. FSH binding to FSH-R stimulated a Protein Kinase A (PKA) dependent,
cyclic AMP response-mediated regulation of a variety of genes, including aromatase (Cyp19A1),
17 β-hydroxysteroid dehydrogenase (Hsd17b) and the LH-choriogonadotropin receptor (Lhcgr).
FSH-R activation was also found to activate the PI3 Kinase pathway in cultured granulosa cells
in a PKA-independent, cAMP-mediated manner exhibited by an increased phosphorylation of
AKT [44] and FOXO1 [45, 46]. Insulin-like Growth Factor-1 (IGF-1) has also been found to be
required for proper antral follicle recruitment as Igf-1 ablation leads to impaired antral follicle
dynamics similar to Fshβ and Fshr deletion [47]. In fact, IGF-1 is believed to enhance FSH
responsiveness in granulosa cells as Igf-1 and Fshr mRNA’s have been found to co-localize in
growing follicles [48]. Moreover, the same study showed that although ablation of Fsh-β did not
change circulating IGF-1 levels, Fshr mRNA was significantly reduced in Igf-1 KO mice.
Similar to FSH signaling, IGF-1 signaling has also been found to activate the PI3 Kinase
11
pathway [49]. Together, FSH and IGF-1 regulation of the Foxo family of transcription factors
(particularly FOXO1), downstream of the PI3 Kinase pathway, is believed to control aspects of
granulosa cell proliferation and survival through Foxo-directed transcriptional modification of a
number of targets: CyclinD2 and p27Kip1, for cell-cycle regulation [46, 50, 51]; Fas ligand, for
regulation of apoptosis [52]; and IGF Binding Protein-1 (IGFBP-1), for a postulated inhibitory
influence on IGF-1 [53].
In addition to the stimulatory effects FSH induces in granulosa cells of antral follicles, basal
levels of LH are able to activate steroid synthesis in the theca cells. As reviewed by Young et al.,
LH-mediated induction occurs in concert with IGF-1 (stimulated by FSH) and KL from
granulosa cells, and a putative impact of GDF-9 from oocytes, that altogether are able to
stimulate theca cell differentiation and steroidogenesis [20]. This leads to the increased
production of androstenedione in the theca which is transported to granulosa cells for conversion
into estradiol via the enzymes aromatase and HSD17β.
Estrogen production is linked with a hormonal negative feedback loop that restricts pituitary
secretion of FSH. This severe reduction in circulating levels of FSH leads to a process of
selection for the recruited antral follicles, leaving only those with highly elevated production of
FSH-R likely to survive. Elevated estrogen levels have alternatively been associated with a
heightened production of LH from the pituitary, which is termed the LH surge. The LH surge
prompts the final phase of follicular ovulation, mediating cumulus granulosa expansion, follicle
rupture, alleviation of the meiotic arrest, release of the cumulus-oocyte complex and formation
of the corpus luteum with the remaining mural granulosa and theca. Meiotic resumption involves
breakdown of the centrally located germinal vesicle (GV), chromatin condensation and
progression through metaphase I (MI) until arrest in metaphase II (MII) awaiting fertilization.
12
1.2.4 Initial and Cyclic Recruitment Factors Associated With Premature
Ovarian Failure
As described above, the total or conditional ablation of various factors involved in ovarian
development, specifically the aspects of initial and cyclic recruitment, can lead to excessive
follicle loss, increased primordial follicle activation, or impaired antral follicle development.
Various genome wide association studies (GWAS), performed on large sample sizes of women
suffering from POF, have independently identified mutations in specific chromosomal regions
associated with POF; and some of these have confirmed the observations of the ablated mouse
models described above. The X-chromosome has been considered a hot-spot for identifying
genes associated with POF, due to excessive follicle atresia noted in Turner Syndrome patients,
in addition to a large number of cases observed with X-chromosome rearrangements [54, 55].
Additional POF mutations noted in human cases include mutations in oocyte factors Bone-
Morphogenetic Protein 15 (BMP15) (located on the X-chromosome), GDF9 and FOXO3, and
granulosa cell factors LHR, FSHR, FOXO1, Estrogen Receptor α (ERα) and CYP19A1 [56-61],
among others not outlined above.
Thus, disruption of either oocyte or granulosa cell factors can lead to impaired folliculogenesis,
depletion of the ovarian reserve, and eventual POF. This implies a dual dependence on the
functionality of both cell types to maintain effective follicular development and survival. This
functionality has been well-documented to be highly reliant on the sustained intercommunication
between both oocyte and granulosa.
13
1.3 OOCYTE-GRANULOSA INTERCOMMUNICATION
Normal oogenesis and folliculogenesis, including the various pathways involved in initial
recruitment, cyclic recruitment and ovulation, are characterized by a great deal of
intercommunication between the oocyte and surrounding somatic cell lineage. One of these
means of cross-talk between oocytes and the surrounding granulosa includes the secretion of
oocyte-derived or granulosa/cumulus-derived factors that can activate receptor-mediated
pathways in the corresponding cell.
1.3.1 Extra-Cellular Signaling
A number of these factors have already been described earlier such as Kit ligand (KL) activation
of oocyte receptor C-KIT [34] and KL and IGF-1 activation of theca cell steroidogenesis [20].
Additionally, paracrine actions of the oocytes have been observed that enable the oocyte to
control granulosa cell-mediated events to regulate proper follicular development. This includes
the oocyte-secreted GDF9 and BMP15 which play important roles in activating and facilitating
granulosa cell proliferation, cumulus cell expansion, cumulus cell differentiation, theca cell
proliferation and cholesterol synthesis [20, 62-64]. Furthermore, oocyte-mediated direct
activation of pathways in the somatic cell lineage has also been linked to the production of
increased metabolite support for oocyte survival [65].
1.3.2 Gap Junction Mediated Signaling
An early report by Anderson et al. described the presence of gap junctions of varying lengths
between the oocyte and its surrounding granulosa cell lineage, forming well before the zona
pellucida, and possibly mediating meiotic progression [66]. Gap junction formation is regulated
14
primarily by Connexin 37 (CX37) and Connexin 43 (CX43). CX37 is found primarily expressed
in gap junctions between oocyte and granulosa cell, whereas CX43 is found in gap junctions
linking granulosa cells [67-70]. These gap junctions are essential for transfer of a number of
metabolites and small molecules directly to the oocyte, from the surrounding granulosa.
Although early follicular development seems unperturbed by the loss of gap junction formation
[70], the presence of these gap junctions are required to maintain proper folliculogenesis,
especially with regards to follicular development, maintenance of meiotic arrest, oocyte survival
and meiotic resumption associated with ovulation.
Cyclic adenosine monophosphate (cAMP) was initially identified as the putative factor involved
in the maintenance of meiotic arrest for oocytes [71]. High levels of cAMP result in the
inhibition of maturation promoting factor (MPF), a complex composed of cyclin-dependent
kinase 1 (Cdk-1/Cdc2) and Cyclin B that mediates aspects of oocyte maturation via cell cycle
control [72]. FSH-stimulated granulosa cells, in addition to G-protein coupled receptors on the
oocyte itself, have been observed to be the source of the oocyte cAMP supply [73-75]. The
cAMP phosphodiesterase (PDE3A) can degrade oocyte levels of cAMP and activate meiotic
resumption [76]. Extensive studies have now implied cyclic guanosine monophosphate (cGMP),
which is produced in granulosa cells and transferred to the oocyte via gap junctions, may be the
key factor that inhibits the activity of PDE3A, preserving the elevated levels of cAMP [77-79].
After recruitment for ovulation, the surge of LH reduces cyclic GMP levels and disrupts gap
junctions between granulosa cells, which results in an overall decrease of cAMP in the oocyte
through active inhibition by PDE3A [77, 80].
15
1.4 OOCYTE METABOLISM
An additional essential role for oocyte-granulosa gap junctions is the supply of metabolites from
somatic cells to the oocyte, further coupling oocyte and granulosa health. Granulosa cells have
been well documented to utilize glucose, pyruvate, lactate and fatty acid breakdown for energy
production; however, the oocyte has revealed an inability to do the same, relying instead on
support from the surrounding granulosa in the form of pyruvate, amino acids, and cholesterol,
transported through the gap junctions [65, 81, 82]. As early as 1967, studies from Biggers et al,
revealed that oocytes that were maintained in a cumulus-oocyte-complex (COC) with cumulus
cell-contact were able to mature in vitro in the presence of lactate, glucose and
phosphoenolpyruvate (PEP) [82]. However, once oocytes were denuded, and removed from
granulosa cell support, in vitro maturation (IVM) only occurred when supplemented with
pyruvate or oxaloacetate, with a very slight increase in maturation rates in the presence of
lactate. No oocyte maturation was observed in the presence of glucose or PEP. These findings
were confirmed by Eppig et al., in 1976, wherein denuded oocytes cultured in the presence of
radioactively labeled metabolites displayed significant utilization of pyruvate with no apparent
breakdown of glucose or lactose [83].
More recent evidence has also indicated that key enzymes involved in glycolysis, amino acid
uptake and cholesterol synthesis are almost completely absent in the oocyte, but highly expressed
in the supporting cumulus cells [84-86]. The small IVM advantage provided by lactate
supplementation was initially thought to be due to lactate conversion into pyruvate within the
oocyte; however, recent studies have implied that it may instead be required for maintenance of
the redox state of the cell [87]. Additional sources of energy for the oocyte include the utilization
of lipid droplet stores which can be broken down during a process known as lipolysis for a
16
supply of fatty acids. These fatty acids are then transported to the mitochondria where they
undergo further breakdown via β-oxidation to provide acetyl coenzyme A (acetyl CoA).
Although COC’s have been shown to contain large quantities of lipid droplets, and undergo an
extensive amount of β-oxidation; the temporal and follicular stage-specific utilization of lipid
storage and breakdown in oocytes has been shown to vary quite drastically between different
species [88]. In mouse oocytes, both in vitro and in vivo studies have revealed that the majority
of lipid synthesis occurs shortly after GV breakdown and meiotic resumption, and hence mature
oocytes and early zygotes have been largely correlated with increased utilization of lipid stores
[89, 90].
Most instances of the previously mentioned studies regarding metabolic uptake have largely been
assayed utilizing COC’s, ovulated oocytes or early zygotes [82, 83, 87, 91]. Recent studies
looking specifically at follicle stage-specific oxidation of pyruvate show that pyruvate uptake
rises with follicular growth, and this is also accompanied by increased oxygen utilization [92].
Pyruvate uptake and oxygen depletion can be utilized as two markers of pyruvate oxidation,
leading to TCA cycle activation and oxidative phosphorylation. Although maximal pyruvate
consumption plateaued after the secondary follicle stage, if normalized for cell volume, primary
oocytes are revealed to be the largest consumer of pyruvate. When this is adapted to oxygen
utilization, breakdown of the pyruvate content of the growing oocyte pool, via TCA cycle and
oxidative phosphorylation, is attributed to be the majority of the energy supply for the oocyte
[92]. Pyruvate dehydrogenase alpha 1 (PDHA1), a subunit of the pyruvate dehydrogenase
complex, is part of a key enzyme complex required for pyruvate breakdown for the TCA cycle.
Oocyte-specific ablation of Pdha1 resulted in a reduction in cellular ATP levels, improper
meiotic maturation, and extensive spindle defects [93].
17
In addition to pyruvate transport through gap junctions, ATP may also be transported across gap
junctions, thus providing a direct energy source for the oocyte [94, 95]. As mentioned
previously, amino acids are also transferred across the gap junctions, and play an important role
in cell maintenance, metabolism and development [81, 84, 96]. They are required for the
production of glutathione, an important reducing agent that is utilized to combat increased
cellular concentrations of reactive oxygen species (ROS) [81]. The amino acid glutamine, in
particular, can be broken down to provide α-ketoglutarate, a substrate of the TCA cycle [97].
Pyruvate and amino acids, aside from a possible direct supply of ATP, have thus been shown to
be essential to provide the oocyte with intermediate metabolites for utilization in the TCA cycle
and for oxidative phosphorylation. This has also been independently supported by studies by
Wycherley et al., highlighting inhibitors of oxidative phosphorylation (cyanide, rotenone, 2,4-
Dinitrophenol) or the TCA cycle (Malonate , Monofluoroacetate) cause disrupted follicular
development and death when used in an in vitro oocyte culture model [98]. Metabolic regulation
of oocyte cell death is itself poorly understood, although various studies on Xenopus egg extracts
have established that the oocyte pentose phosphate pathway maintains inhibition of the cellular
cysteine-aspartic protease Caspase-2 via phosphorylation by the calcium/calmodulin dependant
protein kinase II (CaMKII) [99]. Caspase-2 ablation in mice also affords protection against
primordial follicle death, even upon chemotherapeutic treatment [100], however the mechanistic
pathway and particular factors regulating Caspase activation and resultant oocyte fate, are poorly
understood. Furthermore, GWAS studies, uncovering novel single nucleotide polymorphisms
(SNP) in chromosomal regions of patients with POF, have also demonstrated a role for various
metabolic factors regulating oocyte survival. In particular, they have acknowledged Hexokinase
3 (HK3), required for carbohydrate metabolism; and the mitochondrial DNA polymerase
(POLG) as candidate genes required for maintained ovarian function.
18
Altogether, these studies indicate that the maintenance of oocyte competency, especially prior to
the relief from meiotic arrest associated with maturation, requires a continued dependence on
granulosa cells for an exogenous supply of pyruvate, amino acids and additional metabolites for
energy production via the TCA cycle and oxidative phosphorylation. Disruption of the supply of
nutrients from the granulosa, or the inability of the oocyte to properly metabolize received
nutrients due to impairments in mitochondrial function, can lead to improper folliculogenesis,
defective meiotic maturation, and eventual follicular demise.
1.5 MITOCHONDRIA
The breakdown of pyruvate and amino acids to supply the TCA cycle, and the resultant oxidative
phosphorylation are the fundamental factors of aerobic respiration, and occur at the site of the
mitochondria. Mitochondria have long been considered the ‘powerhouses’ of the cell, supplying
the cell, in this case the oocyte, with an adequate source of ATP for survival. In addition to
energy production, these organelles are the sites of thermoregulation, production of steroid
hormones and are intricately involved in the regulation of cell death.
1.5.1 Mitochondrial Structure and Metabolism
Mitochondria are composed of two membranes, the outer and the inner membrane, the latter of
which surrounds the mitochondrial matrix. Oocyte mitochondria are small and spherical in
shape, with the inner membrane forming convoluted structures termed cristae, around a dense
matrix. Somatic cell mitochondria tend to be longer and more tubular in shape with increasingly
convoluted cristae. The fewer cristae and dense matrix in oocyte mitochondria have resulted in
the belief that these mitochondria are less active than somatic cell mitochondria, and this is
19
supported by the appearance of elongated mitochondria with less dense matrices in the
developing zygote [101]. The inner membrane is the site of the various enzymes involved in the
electron transport chain, whereas the mitochondrial matrix is where the TCA cycle pathway
operates. Pyruvate, fatty acid and amino acid breakdown are delivered through the TCA cycle
resulting in the accompanied reduction of NAD+ and FAD
2+ to NADH and FADH2. NADH and
FADH2 are then fed into the electron transport chain, which is composed of 5 protein complexes
termed Complex I through Complex V. NADH and FADH2 act as electron donors, undergo a
variety of redox reactions, and effectively pump protons across the inner mitochondrial
membrane (IMM). The pumping of these protons creates an electrochemical gradient which is
utilized by Complex V, also known as ATP Synthase. The proton-motive force, driven by the
electrochemical gradient causes the reentry of protons into the mitochondrial matrix, turning the
ATP Synthase motor and synthesizing ATP from ADP. Some electrons can be donated to
molecular oxygen, resulting in the production of reactive oxygen species (ROS). Production of
ROS is believed to occur at the sites of Complex I, II and III [102-105]. High ROS levels have
been implicated in the initiation of various deleterious effects due to oxidative damage,
especially on mitochondrial DNA [106]. However, reduction of ROS levels can occur through
endogenous levels of antioxidants like superoxide dismutases, and glutathione.
1.5.2 Oocyte Mitochondria
Mitochondrial DNA (mtDNA) is a 16kb long, intron-less structure that possesses 37 genes. Of
these 37 genes, 13 code for subunits of the electron transport chain, 2 code for ribosomal RNA’s
and 22 code for transfer RNA’s. The large remaining numbers of subunits for oxidative
phosphorylation, among other mitochondrial processes, are coded by genes in the nucleus. In the
oocyte, each mitochondrion is believed to contain 1-2 copies of mtDNA. MtDNA copy number
20
is believed to increase with oogenesis and folliculogenesis, beginning with a few hundred in
primordial germ cells, and increasing to a few hundred thousand by ovulation [107, 108]. This
sharp increase of the limited mtDNA content from primordial germ cells to the mature oocyte is
termed the mitochondrial bottleneck. Although the mitochondrial bottleneck is believed to result
in a rapid expansion of a small sub-population of mtDNA, thus allowing for the elimination of
mtDNA carrying severe mutations, a number of studies have proposed varying mechanisms and
timelines for this bottleneck [107, 109]. Upon fertilization, mitochondria are maternally
inherited, with paternal mitochondria entering the fertilized zygote, but quickly undergoing
degradation [110, 111].
1.5.3 Mitochondria and Oocyte Developmental Competence
Early studies have suggested that mitochondrial content, specifically mtDNA copy number, can
influence oocyte developmental competence and fertilization [112, 113]. A reduction in ATP
levels has been associated with similar developmental impairments and meiotic spindle defects
in oocytes matured in vitro [114, 115] Induction of mitochondrial damage and increased ROS
have also been shown to decrease ATP levels and give rise to meiotic spindle defects [116-118].
Finally, as mentioned previously, oocyte-specific ablation of pyruvate dehydrogenase, Pdha1
resulted in a reduction in cellular ATP levels, improper meiotic maturation, and extensive
spindle defects [93]. Aging, which has been linked to diminished oocyte quality, has been
associated with a decrease in mitochondrial function accompanied by decreases in ROS and ATP
production [119, 120]. Aged human and mouse embryos have also been associated with
increases in cellular fragmentation [7, 121, 122].
21
Mitochondria are multi-purposed organelles; in addition to regulating aspects of bioenergetics,
they are also the site of the intrinsic pathway of apoptosis. Recent studies, which we will delve
into further ahead, have indicated that mitochondria may be the arena for cross-talk between
areas of metabolic production and decisions of cell fate.
1.6 PROGRAMMED CELL DEATH TYPE 1
In considering oogenesis, folliculogenesis and determinants of oocyte metabolism it is essential
to mention that normal ovarian germ cell loss is extensive during ovarian development and
function. Early estimates have determined that around 99.9% of the total number of germ cells
present in the ovary shortly before birth, do not survive till ovulation [1, 2]. In fact, female germ
cells have been well documented to undergo some form of PCD throughout ovarian
development; specifically during early migration to the gonad, cyst breakdown for primordial
follicle formation, shortly after birth, upon initiation of the growth signal without FSH rescue,
during selection for final ovulation, and other instances. In addition to these factors regulating
natural follicular demise, depletion of the ovarian follicle pool may also occur prematurely.
Untimely depletion of the ovarian follicle pool can be caused by either genetic abnormalities (X-
linked or autosomal recessive mutations), iatrogenic factors (chemotherapy, radiation therapy),
or external environmental exposures (various chemical pollutants), that reduce or abrogate the
starting number of oocytes, or disrupt normal follicle dynamics. As these follicles are eliminated
through some form of PCD, understanding mechanisms that regulate this form of death in
oocytes would allow us to further comprehend and eliminate premature follicular loss.
22
As reviewed by Elmore et al., the apoptotic cascade has been quite well characterized, though
many attributes are still being uncovered [123]. Apoptosis can be divided into two main
pathways: the extrinsic pathway, mediated by death receptors belonging to the Tumor Necrosis
Factor (TNF) family; and the intrinsic pathway, regulated by members of the Bcl-2 family at the
mitochondria. Activation of either pathway does merge upon downstream activation of Cysteinyl
aspartic acid proteases (Caspases), resulting in proteolytic degradation of cell components and
effective apoptotic cell death.
1.6.1 Intrinsic Pathway of Apoptosis
One of the most well-studied gene families responsible for PCD are the anti- and pro-apoptotic
members of the Bcl-2 family. A number of these have been found to have specific functions in
the ovary as well [124]. Bcl-2 was identified using B-cell lymphomas and was described as an
anti-apoptotic oncogene [125]. Additional Bcl-2 members were uncovered due to the presence of
Bcl-2 homology (BH) regions in addition to structural similarities between the members; and
these have been extensively documented [126-128]. The Bcl-2 family is split into core pro- and
anti-apoptotic members, which share several BH domains, and additional pro-apoptotic factors
that carry only a single BH3 domain [128-130]. The core pro-apoptotic Bcl-2 members,
containing multiple BH domain-containing molecules (Bax, Bak, Bok, Bcl2l14/Bcl-G,
Bcl2l13/Bcl-Rambo, Bcl2l15/Bfk), are able to oligomerize, bind and form channels in the
mitochondrial membrane, triggering the apoptotic cascade [127]. The anti-apoptotic members
(Bcl-2, Bcl2l1/Bcl-x, Mcl-1, Bcl2l2/Bcl-w, Bcl2l10, Bcl2l12, Bcl2a1/Bfl1) also possess multiple
BH domains and are believed to function by binding pro-apoptotic members, inhibiting their
oligomerization, and thus neutralizing their killing potential [128]. The core pro- and anti-
23
apoptotic Bcl-2 members also possess a C-terminal Transmembrane (TM) domain that allows the
protein to bind the outer mitochondrial membrane (OMM). Pro-apoptotic BH3-only members
(Bad, Bid, Bcl2l11/Bim, Bik, Bnip3l/Nix,hrk, Bbc3/Puma, Noxa, Bmf, Beclin-1) are structurally
vastly different from the core members, yet the single BH3 domain permits binding to both pro-
or anti-apoptotic members. BH3-only members allow for activation of the death cascade in
response to a variety of stimuli from DNA damage to cellular stress. Current theoretical models
separate these BH3-only members into two groups termed activators (Bim, Bid, Puma) and
sensitizers (Bad, Noxa, Bik, Bmf,hrk, Beclin-1). Activators are believed to directly activate core-
pro-apoptotic members, termed effectors, which bind to the OMM, oligomerize and form
channels in the membrane. Sensitizers, on the other hand, interact solely with anti-apoptotic
members, bind and prevent their activity. Thus far, a variety of models have been proposed
attempting to describe the actual mechanisms of activation/inhibition of the effectors and the
eventual stimulation of the death cascade; however none have been fully validated.
Additionally, different activators or sensitizers have shown varying binding affinities for pro- or
anti-apoptotic Bcl-2 members. Binding of BIM, BID or PUMA to BAX and BID to BAK has
been demonstrated in in vitro studies [131-134], and a triple knockout of Bid/Bim/Puma
displayed near replication of the Bax/Bak double knockout phenotype [135]. For BH3-only
binding to anti-apoptotic members, an in vitro analysis was performed by Chen et al, in order to
determine a number of these binding affinities [136]. BIM and PUMA were found to effectively
bind all anti-apoptotic members, whereas BID, BIK, andhrK showed strong selective binding for
BCL-2, BCL-xL and BFL-1. BAD and BMF showed preferential binding with BCL-2, BCL-xL
and BCL-w, and NOXA was found to restrictively bind to MCL-1 and BFL-1. Additional in
vitro studies have enlarged this model which have been collectively displayed in a notable
review by AJ Garcia-Saez [137].
24
Stimulation of the intrinsic pathway through activation of a death signal results in the
oligomerization of the pro-apoptotic effector Bcl-2 family members and the formation of a pore
in the OMM. Pore formation allows the release of a number of factors from the mitochondrial
intermembrane space into the cytosol, which activates the apoptotic cascade [138] (Fig. 1.3). The
actions of these various factors have been studied extensively and, as reviewed in Elmore et al.
[123], are composed of Cytochrome c, Second mitochondrial activator of caspases
(SMAC)/Direct IAP binding protein with low PI (DIABLO), High-temperature requirement
protein A2 (HTRA2)/Omi, Apoptosis Inducing Factor (AIF), Endonuclease G and Caspase-
Activated DNAse (CAD). Cytochrome c (CYC1) binds to the Apoptotic protease activating
factor (APAF-1) and pro-caspase 9 (forming the apoptosome), then cleaves and activates the
initiator caspase, caspase-9. Active SMAC/DIABLO is able to assist caspase cleavage by
antagonizing the action of Inhibitor of Apoptosis Proteins (IAPs). HTRA2/Omi is a serine
protease that binds and cleaves IAPs. AIF has been shown to relocate to the nucleus upon
induction of cell death and leads to chromatin condensation and DNA fragmentation. Recent
evidence has shown that AIF can also be localized to the nucleus in a caspase-independent
manner, and may be linked to necrotic cell death [139], or an independent AIF-mediated PCD
[140]. Endonuclease G is a DNAse that is also directed to the nucleus and participates in DNA
fragmentation. Furthermore, like AIF, it has also been shown to activate DNA fragmentation in a
caspase-independent manner [141]. CAD is a caspase activated DNAse, that is cleaved and
activated by the executioner caspase, Caspase-3 (CASP-3), and translocates to the nucleus
resulting in further DNA fragmentation and more intense chromatin condensation. Executioner
caspases are cleaved and activated by the initiator caspases, and direct the proteolytic destruction
of the cell.
25
Figure 1.3. Intrisic Pathway of Apoptosis.
Bcl-2 pro- and anti-apoptotic members mediate the intrinsic pathway of apoptosis. BH3-only sensitizers
(Bad, Noxa, Bik) inhibit the activity of the anti-apoptotic Bcl-2 members (Bcl-2, Bcl-x, Mcl-1),
preventing their inhibition of Bcl-2 pro-apoptotic effector proteins (Bax, Bak, Bok) or BH3-only
activators (Bim, Bid, Puma). When Bcl-2 anti-apoptotic inhibition is relieved, BH3-only activators can
activate effector proteins which bind to the mitochondrial membrane, oligomerize and form a pore in the
outer mitochondrial membrane (OMM). This allows the release of a number of factors from the inter-
membrane space (IMS) into the cytoplasm, which mediate cellular destruction. These factors include:
AIF, Endonuclease G and CAD, which are transported to the nucleus and direct DNA fragmentation and
chromatin condensation; Smac/Diablo and HtrA2/Omi, which impede the action of Inhibitor of Apoptosis
Proteins (IAP’s); and Cytochrome C, which binds Pro-Caspase 9 and APAF-1 to from the Apoptosome,
resulting in the cleavage of pro-Caspase 9 to form the active initiator caspase, Caspase 9. Caspase 9 then
cleaves pro-Caspase 3 activating the executioner caspase Caspase 3, which mediates the proteolytic
degradation of cellular components.
26
1.7 PROGRAMMED CELL DEATH TYPE 1 IN THE OVARY
Oocytes have been documented to undergo various means of cell death in a stage- or
environmentally-specific manner. Assessments of the in vivo growing follicle pool have
determined that follicular death can occur with expression of both autophagic and apoptotic
factors, and markers of fetal oocytes have indicated their ability to activate various modes of cell
death [142-144]. New studies are challenging the classical notion, indicating that primordial
follicles in the postnatal ovary do not undergo a classical apoptotic death [145]. These studies
imply that in the postnatal prepubertal ovary, the lack of classical apoptosis is due to
maintenance of granulosa and oocyte contact, following primordial follicle formation. Notably,
follicular atresia of growing follicles has been documented as a granulosa cell-mediated event,
initiated due to granulosa cell death [146]. It is characterized by granulosa cell withdrawal from
the oocyte, severing oocyte contact and trans-zonal projections [147]. Ovulated oocytes undergo
fragmentation, which has been classified as containing hallmarks of apoptotic cell death [148]
1.7.1 Pro-apoptotic Bcl-2 Family in the Ovary
Various Bcl-2 pro- and anti-apoptotic members have been studied to determine differing patterns
of expression and functional importance in oogenesis and folliculogenesis. Pro-apoptotic BAX
was initially identified in granulosa cells, and Bax mRNA levels were reduced upon
gonadotropin stimulation, also associated with follicular survival [149]. Bax-deficiency was also
initially associated with male infertility, and an accumulation of granulosa cells in atretic
follicles [150], indicative of impairment in granulosa cell apoptosis. Bax mRNA was also
identified in GV oocytes [151], and Bax ablation was shown to provide protection against
27
chemotoxic-induced primordial and primary follicle death [152] and chemotherapy-induced
death in primordial follicles and mature ovulated oocytes [153, 154]. Bax ablation also provided
protection against Polycyclic Aromatic Hydrocarbon (PAH)-induced primordial follicle death
[155]. Deletion of Bax was linked to an extension of fertility with the alleviation of some age-
related health issues [156, 157]. This was associated with the persistence of primordial follicles
and a reduction in atretic follicles in young females, and the maintenance of healthy follicles in
aged Bax-deficient mice. These studies indicated that the elongation of ovarian function was due
to reduced postnatal follicle atresia, and this was supported by no apparent change in the
primordial follicle pool number, shortly after birth. Additional studies have shown that Bax also
plays a role in germ cell fate during early germ cell migration and population of the fetal gonad
[158, 159]. These, in addition to results indicating that embryonic and neo-natal Bax-deficient
females do have an increase in germ cell number [160], imply that the elongation of fertility may
be a result of an augmented primordial follicle pool endowment due to increased germ cell
survival during migration. Furthermore, recent evidence has also determined that Bax may be
inessential for follicular atresia [161].
BAK expression was found in the thecal cells of the human ovary [162], however deletion of
Bak was dispensable for both male and female gonadal development and function [163].
Expression of the pro-apoptotic member Bok mRNA was localized to granulosa cells of preantral
and antral follicles [164]. BOK protein expression was identified in the nucleus and cytoplasm of
oocytes of fetal human ovaries, and in oocytes and granulosa cells of growing follicles in adult
human ovaries [165]. Jaaskelainen et al. also displayed that Bok ablation was able to protect
granulosa cells in vitro in the presence of apoptosis-inducing factors. Although BOK is strongly
expressed in reproductive tissues, its ablation had no impact on reproductive development or
function [166].
28
Granulosa cells of all follicle stages in the postnatal ovary were found to express the pro-
apoptotic BAD [167]. Overexpression of BAD in primary granulosa cell lines activated
apoptosis in a caspase-dependant manner. PAH exposure induced pro-apoptotichrK expression
in oocytes and granulosa of primordial and primary follicles in neonatal mice [168]. PAH-
induced primordial and primary follicle death was demonstrated to occur viahrK-induced BAX
activation. Furthermore, BID was identified as an additional pro-apoptotic factor that interacted
with BAX to regulate granulosa cell death during follicular atresia [169, 170].
mRNA expression of Bim, also known as Bcl-2-related ovarian death gene (BOD), was noted in
ovaries utilizing Northern Blot techniques [171]. Bim deletion resulted in a variety of
lymphocytic defects, however no fertility issues were observed [172]. Yet, expression of two of
the three known Bim isoforms, Bim-long (BimL) and Bim extra-long (BimEL), were detected in
granulosa cells of all stage follicles in mice and pig, and primordial follicle oocytes in rat ovaries
[173-175]. Additionally, induction of BIM was associated with oocyte and granulosa cell
apoptosis, and was transcriptionally regulated by the PI3Kinase pathway, specifically by the
Foxo transcription factors [173].
Puma and Noxa expression was noted in oocytes of primordial follicles in ovaries of postnatal
day 5 mice exposed to γ-irradiation (0.45-4.5 Gy) [176]. γ-irradiation also resulted in an
induction of PUMA protein expression. Interestingly, this induction was not present in the
oocytes of TAp63-deficient mice. TAp63, a transcription factor and member of the Trp53 family
of tumor suppressors, has been documented with initiation of DNA damage-induced primordial
oocyte apoptosis in in vivo mouse models [177, 178]. Furthermore, Kerr et al demonstrated that
Puma-deficient females were afforded a greater protection against primordial follicle death via
radiation-induced DNA damage, and remained fertile; and this protection was enhanced with the
29
additional ablation of Noxa. However, Noxa-deficiency alone did not prevent primordial follicle
loss.
1.7.2 Anti-apoptotic Bcl-2 Family in the Ovary
Initial studies using Bcl-2 deficient mouse models revealed that Bcl-2 plays a pro-survival role in
the ovary, as ablation of Bcl-2 led to a small but significant reduction in primordial follicles at
PN42 [179]. Transgenic overexpression of Bcl-2, under the control of the C-Kit promoter,
resulted in an increased primordial follicle pool shortly after birth (PN8); however this increase
did not persist 1-2 months postnatally [180]. In order to determine the contribution of BCL-2 to
the primordial follicle pool at birth, Jones et al. analyzed Bcl-2-deficient mice and Bcl-2
overexpressing transgenic mice under the control of the C-Kit promoter [181]. They found no
change in primordial follicle numbers at birth, and hence attributed the change in primordial
follicle number found in the previous studies to be a result of a postnatal impact of BCL-2 on
primordial follicle survival. Additionally, BCL-2 expression has been localized to granulosa cells
of growing follicles and corpura lutea in the rat ovary, and may play a role in mediation of cell
survival therein [182]. Overexpression of Bcl-2 in granulosa cells resulted in increased granulosa
cell survival, increased ovulation rates implying increases in follicular development, and
increased germ cell tumor formation [183].
BCL-x expression was originally noted in oocytes and granulosa cells, in addition to corpora
lutea in the post-pubertal mouse ovary [184]. Total ablation of Bcl-x resulted in embryonic loss
as early as embryonic day 12.5 [185]. To circumvent the embryonic lethality, Rucker et al.
created a hypomorphic Bcl-x allele by inserting a Neomycin resistance (Neo) cassette within the
30
Bcl-x promoter region flanked by two loxP sites [158]. The presence of two hypomorphic Bcl-x
alleles led to embryonic disruption and reduction in the number of fetal germ cells that populated
the embryonic gonad; leading to a severe impairment in primordial follicle number in postnatal
ovaries. Removal of the hypomorphic alleles using Cre Recombinase-mediated excision restored
germ cell number to levels comparable to wildtype. Bax deletion in the same mice was able to
rescue this phenotype, thus indicating a potential BAX/BCL-x interplay that regulates fetal germ
cell fate. In the adult hen, BCL-x was identified in granulosa cells of growing follicles, and an
increase in BCL-x expression was associated with an increase in granulosa cell survival [186].
However, postnatal conditional ablation of Bcl-x in oocytes, granulosa cells, and luteal cells
revealed no apparent requirement for Bcl-x [187]. These findings indicate that Bcl-x, although
required for primordial germ cell survival in male and female gonads, is relatively inessential in
postnatal follicular fate.
Diva, also known as Boo, BCL2-like 10 (BCL2L10) or Bcl-B, was localized to the granulosa
cells of growing follicles [188]. Diva expression was also found in immature murine and human
GV oocytes and was maintained into the oocyte-zygotic transition [189]. Total ablation of Diva
resulted in no apparent ovarian phenotype, and provided no protection against radiation-induced
genotoxic stress of the ovary or other tissues [190].
The anti-apoptotic function of Bcl-w was found to be specific for mediating spermatogenesis in
adult testis [191]. No functional role for Bcl-w in ovarian development was ascertained. These
data imply that either the postnatal functional role of Bcl-x and Bcl-w in normal ovarian
development does not exist; or that it may prove to be redundant, fulfilled by other anti-apoptotic
Bcl-2 members. However, thus far, no anti-apoptotic Bcl-2 member has been attributed to a role
31
in governing postnatal oocyte survival, although a mildly protective role has been ascribed to
Bcl-2.
Limited work has been performed on the role of the anti-apoptotic member Myeloid Cell
Leukemia 1 (MCL-1) in folliculogenesis. Early work has established expression in fetal human
oocytes of the primordial follicle pool, murine oocytes of preantral follicles and granulosa and
theca cells of growing follicles; and has implicated Mcl-1 with a role in primordial follicle
survival [181, 192, 193]. The role of Mcl-1 in the ovary will be studied in further detail in the
upcoming chapters.
1.8 PROGRAMMED CELL DEATH TYPE 2
PCD Type 2, or autophagic cell death, has been revealed as another means by which a cell, in the
absence of proper nutritional sustenance, turns to self-digestion to provide itself with energy. At
first considered a means of cell death, it has also been displayed that autophagy can occur to
eliminate malfunctioning organelles and provide energy in times of stress, and thus can also be
referred to as a cell survival mechanism. The mechanistic process of autophagy has been well
documented in a large number of reviews [194-196].
Macroautophagy is the process via which the cell degrades various cellular components in times
of stress or starvation, in order to maintain a steady supply of ATP for cell survival. In this thesis
I shall be referring to all cases of macroautophagy as simply autophagy. Briefly, autophagy
begins with the formation of an autophagosome which surrounds organelles that have been
designated for degradation; the autophagosome then fuses with a lysosome to form an
autolysosome, and the substrates within are degraded to release nutrients for the cell (Fig. 1.4).
32
33
Figure 1.4. Molecular Pathways of Autophagy.
In times of nutrient starvation, mTORc1 kinase repression of ULK complex activity is alleviated and a
disrupted ATP/AMP ratio results in activation of AMP Kinase activity. ULK complex and AMP Kinase
activity leads to phosphorylation and stimulation of Beclin-1 and Class III PI3 Kinase VPS34, and
recruitment to the site of autophagosome formation. The BH3 domain located on Beclin-1 allows for
binding and inhibition of autophagy by Bcl-2 anti-apoptotic members. Activated Beclin-1 complexes with
VPS34 and ATG14L to form the VPS34 complex and phosphorylates phosphatidylinositol to form PI3P.
Through recruitment and action of various Autophagy related (Atg) genes, PI3P and VPS34 complex
recruits and induces formation of the ATG16L complex, formed of ATG12, ATG5 and ATG16. The
ATG16L complex mediates conjugation of the lipid PE to LC3, which together with lipids acquired from
mitochondrial or ER source begins development of the isolation membrane. Adaptor proteins with LC3-
interacting regions bind substrates for degradation and recruit them to the site of autophagosome
formation. After formation is complete, numerous lysosomes containing lysosomal hydrolases merge with
the created autophagosome forming an autolysosome, and degrade the internal contents. These are then
released into the cytoplasm and the lysosomes are reconstructed.
34
1.8.1 Autophagosome Formation
The formation of the autophagosome is preceded by the induction of autophagy, which is held in
check by the activity of the mTOR Complex1 [197]. During conditions of starvation, mTORc1
kinase activity is impaired, and thus it is unable to complex with, phosphorylate, and inhibit the
activity of the Autophagy related 1 (ATG1)/Unc51-like kinase (ULK) complex [198].
Downstream factors phosphorylated by ULK complex activity are important elements in
autophagosome formation, and recruitment of ATG proteins to the formation site. Additionally,
recent studies have shown that, during amino acid starvation, ULK is also essential for
phosphorylation of Beclin-1(BECN-1) and resultant activation of the Class III PI3 Kinase VPS34
[199]. VPS34 is able to phosphorylate phosphotidylinositol to form phosphatidylinositol 3-
phosphate (PI3P), which is required for autophagosome initiation. Furthermore, new evidence
has determined that Adenosine-monophosphate protein activated-kinase (AMPK), which is
uniquely sensitive to starvation (low ATP/high AMP) conditions [200], can directly
phosphorylate either Beclin-1, VPS34 or ULK directly to initiate autophagosome formation
[201, 202]. Anti-apoptotic Bcl-2 family members have also been demonstrated to inhibit
autophagic activation by binding to the BH3 domain located on Beclin-1 [203-205].
Autophagosome initiation begins with the formation of an isolation membrane. Although the
Endoplasmic Reticulum was initially considered to be the major source of membrane for the
autophagosomes, recent studies have displayed that during starvation-induced autophagy, the
outer membrane of the mitochondria provides a majority of the membrane structure [206].
Formation and elongation of this isolation membrane is an intricate process involving lipid
recruitment, complex formation and conjugation of ATG5, ATG12, and ATG16, in addition to
conjugation of the lipid phosphotidylethanolamine (PE) to the Microtubule-associated protein 1
35
light chain (MAP1LC3A) also known as simply LC3 [196]. Conjugation of PE to LC3 results in
an autophagic membrane bound conformation of LC3, referred to as LC3II. The complex formed
by ATG5, ATG12 and ATG16, known as the ATG16L complex, is believed to be required for
LC3 transport to the autophagosome formation site [207]. Although once considered a
macroscopic degradation of numerous organelles; new studies, as summarized by Johansen et al.,
indicate that there are adaptor proteins allowing for selective targeting of various substrates to
the autophagosome [208]. These adaptor proteins can contain LC3-interacting regions (LIR) that
direct recruited substrates to the site of autophagosome formation. Recent evidence, as put
forward by Morita et al., indicates that LC3 might not be required for formation of the isolation
membrane, but might just be utilized for substrate-targeting [209]. However, they do
acknowledge the requirement for LC3 in autophagosomal membrane closure [210].
1.8.2 Formation of Autolysosome and Substrate Degradation
After formation, the autophagosome can fuse with multiple lysosomes to form an autolysosome,
or an autophagolysosome. This fusion leads to degradation of the contents of the autophagosome
by lysosomal hydrolases. Lysosome-associated membrane proteins 1 and 2 (LAMP-1, LAMP-2)
are an integral part of the autophagosome maturation. These membrane-bound proteins have
been found to be essential for autolysosome formation and lysosomal fusion, possibly via their
ability to mediate transport through interactions with the microtubular motor complex of the cell
[211]. After degradation of the autolysosome contents, lysosomes are reconstructed through the
formation of autolysosomal buds that give rise to LAMP-positive, but LC3-negative tubular
membrane structures that form proto-lysosomes [212]. These proto-lysosomes then mature to
form functional lysosomes.
36
1.9 AUTOPHAGY (PCD TYPE 2) IN THE OVARY
Recent studies have identified the activation of the autophagic pathway in oocytes [142].
Extremely limited work has been performed on the induction and utilization of autophagy in
oocytes, and although it has been found to be required for early zygotic development [213],
factors that regulate its induction in the growing follicle oocyte pool, and follicle atresia, are not
fully understood. Analyses of markers of autophagy in oocytes have revealed that follicles
undergoing atresia express LAMP1 and acid phosphatase, which are markers of lysosomes, in
addition to the ultrastructural appearances indicative of autophagosome formation. Markers of
apoptosis were concurrently expressed. An additional study utilizing follicles from different
estrous stages also observed cytoplasmic autophagosome formation in addition to increased
intensity in expression of LAMP1 and LC3 in those follicles that were undergoing atresia [214].
Furthermore, differing levels of LC3 and LAMP1 were noted in various oocytes, indicating basal
levels of autophagy.
Various models utilizing ablation of autophagy components have been created in the ovary.
Beclin-1, required for initiation of autophagosome formation, was initially localized to theca
cells, in addition to granulosa and theca cells undergoing luteinization [215]. Further studies
displayed that Beclin-1 mRNA was noted in granulosa, theca and oocytes of all stage follicles,
with strong expression noted in primordial follicles [216]. Becn-1 heterozygote females
displayed a significant reduction in the endowment of the primordial follicle pool, shortly after
birth. The same study also revealed that mRNA expression of Atg7, involved in isolation
membrane elongation, was observed in oocytes of all follicle stages. When ablated, Atg7 also
resulted in a severe depletion in the primordial follicle pool.
37
1.9.1 Bcl-2 Family in Autophagy and Metabolism
The Bcl-2 family has recently been conferred with roles in not just regulation of apoptosis, but
additional functions in autophagy and metabolism. Anderson et al. have reviewed the various
factors linking metabolism and apoptosis, and specifically the overlapping role of the Bcl-2
family [217]. The role of the Bcl-2 family in autophagy has also been studied in some detail.
Initial observations that Bcl-2 family members were found to bind to Beclin-1 in neurons, were
shortly followed by inferences that BCL-2 binding to Beclin-1 was required for inhibition of
Beclin-induced autophagy [205, 218]. Beclin-1 was also established to bind Bcl-2 anti-apoptotic
members BCL-x, BCL-w and MCL-1 in mammalian cell lines, utilizing a BH3-like domain;
binding to pro-apoptotic members was not observed [204, 219].
Bcl-x and Mcl-1 have both been associated with particular roles regulating oxidative
phosphorylation. BCL-xL has been implicated in prevention of proton leakage and increased
ATP Synthase pump efficiency in neurons [220]. A mitochondrial matrix-specific isoform of
MCL-1 has been associated with enhanced ATP Synthase assembly in mouse embryonic
fibroblasts (MEFs), a function found to be distinct from its anti-apoptotic role [221]. Increased
glucose metabolism has also been demonstrated to increase MCL-1 stability, through inhibitory
regulation of GSK-3 in a PI3 Kinase pathway dependent manner [222, 223]. This relationship
will be studied in greater detail shortly.
Mcl-1, like other Bcl-2 family members, has been found to be an integral factor involved in
apoptosis, cell metabolism and autophagy, and we shall analyze this impact in greater detail
ahead.
38
1.10 MCL-1
MCL-1, an anti-apoptotic Bcl-2 family member identified in early human myeloid leukemia cell
lines (ML-1), was observed to be induced early in ML-1 cells that were undergoing
differentiation [224]. MCL-1 has a high degree of sequence similarity to BCL-2 and induction of
either was related to a survival-based mechanism without increased proliferation. Further studies
determined that MCL-1 is widely expressed in many tissues, and Mcl-1-deficiency in mice leads
to peri-implantational lethality [225]. Mcl-1-deficient embryos can be recovered at 3.5-4dpc yet
exhibit an implantation defect due to the failure to form a trophoectoderm outgrowth. However, a
unique feature of the Mcl-1-deficient embryos isolated by Rinkenberger et al., was the absence of
apoptosis in the blastocyst, and the lack of rescue with concurrent Bax or p53 deficiency. This
indicated that there may be additional roles for Mcl-1 besides the classical apoptotic angle.
Utilizing a Cre-lox inducible system for cell-specific Mcl-1 deletion, Mcl-1 was observed to be
essential for hematopoietic stem cell survival in addition to B and T lymphocyte survival [226,
227]. Mcl-1-conditional ablation in neutrophils led to increased neutrophil apoptosis, which was
rescued by concurrent Bax and Bak deletion; whereas conditional Mcl-1-deletion in macrophages
resulted in an increased sensitivity to cell death, and was rescued by simultaneous deletion of the
BH3-only activator Bim [228, 229]. Finally, recent work by Wang et al., has revealed that Mcl-1-
deficient cardiomyocyctes have severe cardiac defects, along with a variety of mitochondrial
structural and respiratory dysfunctions [230]. Simultaneous deletion of Bax and Bak rescued a
number of the Mcl-1-deficient cardiac phenotypes, however mitochondrial function remained
impaired. Additional work utilizing the Cre-lox system for Mcl-1 deletion in fibroblasts, has also
led to the postulation of new roles of Mcl-1 in mitochondrial matrix-directed mitochondrial
respiration [221].
39
1.10.1 Gene, Transcript and Protein Structure
Mcl-1 is a 3 exon gene giving rise to a 331 amino acid protein in mouse (Human-350aa) that is
considerably larger than its other anti-apoptotic Bcl-2 counterparts (Fig. 1.5). This is due to the
large N-terminal PEST domain, composed of Proline, Glutamic Acid, Serine and Threonine
residues, that leads to MCL-1 being quite susceptible to a number of post-translational
modifications. These modifications have the ability to mediate various aspects of MCL-1
stabilization and degradation [231, 232]. MCL-1 is characterized by 3 BH domains allowing it to
bind to other members of the Bcl-2 family, and a TM domain, permitting OMM permeability.
An alternatively spliced isoform of MCL-1, termed Mcl-1 short (MCL-1S), was found in human
placenta [233]. This alternative splicing results in the removal of exon2, a frameshift, and the
subsequent loss of the BH1, BH2 and TM domains, but the maintenance of the BH3 domain. An
additional splicing variant, Mcl-1 extra short (MCL-1ES) was revealed in human cell lines, and
was demonstrated to lack a significant portion of exon 1 resulting in the loss of the PEST domain
[234]. However, MCL-1ES sequences have been shown to retain all three BH domains in
addition to the TM domain. Both splicing isoforms have been demonstrated to be pro-apoptotic,
with proposed activities including binding and inhibition of full length MCL-1 (Fig. 1.5).
1.10.2 Transcriptional Regulation of Mcl-1
A number of factors have been identified that regulate Mcl-1 transcription, and these have been
reviewed by Thomas et al [231]. A majority of cytokines have been demonstrated to increase
Mcl-1 transcription and these include epidermal growth factor (EGF), vascular endothelial
growth factor (VEGF), Interleukin 3 (IL-3), IL-5, IL-6, KL and granulocyte-macrophage colony-
stimulating factor (GM-CSF) [231, 235]. Transcription factors directly associated with increases
40
in Mcl-1 expression include activating transcription factor 5 (ATF5), signal transducer and
activator of transcription 3 (STAT3), PU box binding transcription factor (PU.1), cyclic AMP
response element binding protein (CREB), Specificity protein (SP-1), hypoxia-inducible factor
1α (HIF-1α) and nuclear factor κ B (NFκB); whereas E2F transcription factor (E2F-1) decreases
Mcl-1 transcription [231, 232].
1.10.3 Post-Translational Regulation of MCL-1
MCL-1 has been documented to have an extremely short half-life, with turnover rates being
anywhere from a few hours, to as short as 30 minutes, depending on treatment and cell lineage
[236]. Proteosomal degradation of MCL-1 was initially correlated with increases in apoptosis
upon ultraviolet exposure of human cell lines [237]. This occurs via phosphorylation of MCL-1
at particular residues which increases recruitment of ubiquitin ligases that ubiquitinate MCL-1
and lead to increased proteosomal degradation (Fig. 1.5). Several ubiquitin ligases have been
identified that regulate MCL-1 stability including HECT, UBA and WWE domain containing-
(HUWE1)/Mcl-1 ubiquitin ligase E3 (MULE), β-transducing repeat containing E3 ubiquitin
ligase (β-TrCP), and F-box/WD repeat domain (FBW7) containing ligase subunit of Skp1, Cul1
and F-box protein (SCF) ubiquitin ligase complex [238-240]. In order to mediate excessive
action of these various enzymes, a de-ubiquitinase, termed ubiquitin-specific peptidase 9 X-
linked (USP9X) has been identified that removes ubiquitin residues hence prolonging MCL-1
stability [241]. An additional arm of complexity was established in studies by Gomez-Bougie et
al., where the pro-apoptotic BH3-only sensitizer NOXA was found to further regulate the MCL-
1/USP9X/MULE axis [242]. Induction of NOXA was found to de-stabilize the MCL-1-USP9X
complex, increasing MCL-1 polyubiquitination and escalating the MCL-1-MULE interaction;
thus resulting in elevated MCL-1 degradation.
41
Furthermore, MCL-1 degradation has also been associated with caspase-dependent cleavage at
Asp127 and 157
residues within the N-terminal PEST domain [243, 244]. TRAIL, a member of the
extrinsic pathway of apoptosis, was only able to induce apoptotic cell death in human cell lines
when accompanied by caspase-dependent cleavage of MCL-1, permitting reduction of MCL-1
inhibition of pro-apoptotic Bcl-2 family members, specifically BIM [244, 245]. MCL-1
degradation via Granzyme B-mediated cleavage was also identified; occurring at Asp117
, Asp127
and Asp157
and also instigating disrupted MCL-1 sequestration of BIM, resulting in increased
apoptotic potential [246, 247].
The large number of residues in the PEST domain allows for MCL-1 regulation through a variety
of pathways. Phosphorylation at the Serine 64 residue has been attributed to activity of cyclin
dependent kinases (CDK1 and CDK2) and cJun N-terminal kinases (JNK1) [248, 249]. Although
phosphorylation at this residue does not modify MCL-1 turnover rates, it has been shown to be
cell-cycle dependent, with the greatest phosphorylation occurring at the G2/M phase. Ser64
phosphorylation is believed to modify binding efficiency of MCL-1 to pro- and anti-apoptotic
members of the Bcl-2 family, and thus regulate sensitivity to apoptotic triggers.
Phosphorylation by extracellular signal-regulated kinases (ERK), members of the well-known
mitogen-activated protein kinase (MAPK) pathway, at the Threonine 163 (Thr163
) residue was
found to increase MCL-1 stability [250]. Thr163
is categorized as a ‘priming’ residue, required to
be phosphorylated in conjunction with other residues, for the effective increased or reduced
stability/anti-apoptotic activity of the MCL-1 protein. ERK-mediated phosphorylation at Thr92
and Thr163
has been documented to increase MCL-1 stability [251];
42
Figure 1.5. Mcl-1 mRNA Transcript and Protein Structure.
Mcl-1 is a 3-exon gene that gives rise to a 331aa protein in mouse (350aa human). There are three
documented isoforms of Mcl-1: the full length; the splice variant termed Mcl-1-Short (Mcl-1S) isoform,
identified in human placental lines formed by skipping of exon 2, that leads to production of a 271aa
protein; and the splice variant termed Mcl-1-Extra Short (Mcl-1ES), identified in a variety of human cell
lines, that encodes a 197aa protein due to a truncation in exon 1. The MCL-1 protein is characterized by a
long N-terminal PEST domain, susceptible to numerous post-translational modifications; in addition to a
BH1, BH2 and BH3 domain, that allows binding with other Bcl-2 members, and a large trans-membrane
(TM) region, which permits binding to the outer mitochondrial membrane. In addition to two caspase
cleavage sites, the full length MCL-1 protein also has an N-terminal mitochondrial processing peptidase
cleavage site that results in internalization and mitochondrial matrix localization of the cleavage product.
Formation of MCL-1S results in a frame-shift and loss of BH1, BH2 and TM region, whereas MCL-1ES
retains all domains but loses a large chunk of the PEST domain.
43
Phosphorylation of Serine 121 (Ser121
) has also been determined to occur in combination with
Thr163
via activation of the JNK Kinase and p38, members of the MAP Kinase pathway [252,
253]. Initial studies demonstrated that JNK Kinase activity was required to phosphorylate and
inhibit MCL-1 activity in response to hydrogen peroxide treatment [252]. More recent studies
using TNF-induced hepatocyte apoptosis have determined that MCL-1 phosphorylation at Ser121
and Thr163
by JNK kinase actually stabilizes MCL-1, reducing its turnover rate and thereby
increasing its anti-apoptotic potential [253].
The Ser155
residue was found to be phosphorylated in concert with Ser159
and Thr163
by the GSK-
3[238]. GSK-3 phosphorylation of MCL-1 was followed by ubiquitin ligase recruitment and
subsequent degradation of MCL-1. This degradation was circumvented by mutation of the three
phosphorylation sites. Maurer et al., demonstrated that phosphorylation of MCL-1 at Ser159
resulted in the reduced stability of MCL-1, its increased turnover rate, and a reduction in binding
to the pro-apoptotic BIM [222]. Additionally, JNK kinase phosphorylation of the Thr163
residue
has been shown to be required to ‘prime’ this GSK-3-mediated phosphorylation of Ser159
upon
UV-irradiation exposure [254].
1.10.4 Metabolic Role of MCL-1
The PI3Kinase pathway, MAP Kinase pathway and the mTOR pathway have all been
documented to regulate GSK-3 activity, downstream of a multitude of growth factors. Thus,
GSK-3 phosphorylation of MCL-1 may place MCL-1 downstream of a variety of growth factor
induced signaling pathways. In addition, GSK-3 phosphorylation of MCL-1 has been
documented to be decreased following increased glucose metabolism, due to upstream inhibition
44
by the PI3Kinase pathway, resulting in increased MCL-1 protein stability [222, 223].
Additionally, studies by Coloff et al. demonstrated that glucose deprivation, glycolytic
inhibition, or growth factor withdrawal of mouse cell lines resulted in a reduction of MCL-1
levels [255]. Interestingly, constitutively active AKT kinase-dependent rescue of MCL-1
expression levels was reliant on the presence of glucose, and the authors attribute this to
mTORc1 and AMP Kinase regulated pathways.
An additional post-translational cleavage isoform of MCL-1, termed the fast-moving (FM) or
MCL-1 matrix-localized isoform, has been identified as an integral component of the
mitochondrial matrix [221, 256]. Formed via N-terminal cleavage by mitochondrial processing
peptidase (MPP) (Fig. 1.5), it has been associated with enhanced ATP Synthase assembly in
MEF’s, a function found to be distinct from its anti-apoptotic role [221]. This provides for an
additional putative role of MCL-1 in the maintenance of cell metabolism and regulation of
mitochondrial bioenergetics.
1.10.5 Autophagic Role of MCL-1
Mcl-1 has been shown to play an avid role in regulation of autophagy, in addition to its
metabolic and anti-apoptotic role. As mentioned earlier, Beclin-1, required for initiation of
autophagosome formation, was discovered to bind MCL-1 in Glutathione S-transferase (GST)-
tagged pull down assays in human cell lines [219]. Furthermore, mutation of the BH3 domain in
Beclin-1 disrupted interaction with MCL-1; and MCL-1 expression was also able to inhibit the
induction of autophagy due to overexpression of Beclin-1 [204]. Additional work by Germain et
al., confirmed the interaction of MCL-1 and Beclin-1, in addition to determining that under
45
nutrient deprivation conditions, degradation of MCL-1 preceded autophagosome initiation, and
overexpression of MCL-1 prevented autophagic initiation [203]. In fact, ablation of floxed MCL-
1 in cortical neurons in vitro with exogenous Cre, in addition to in vivo ablation in various
neurons using a Ca2+/calmodulin-dependent protein kinase (CamKIIα)-promoter driven Cre,
revealed increased activation of the autophagic pathway. Utilizing further in vitro and in vivo
analyses, Germain et al. revealed that deletion of MCL-1 in cortical neurons resulted in the
activation of autophagy, with a subset eventually activating apoptosis. These authors surmised
that down-regulation of MCL-1 potentiated the cell for activation of either autophagy or
apoptosis, dependent on the cellular stress or context of the cell. Additionally, autophagic
responses have been displayed to lead to apoptosis by either activation of the apoptotic triggers,
inhibition of the autophagic pathway, or caspase-mediated cleavage of Beclin-1 [203, 257, 258].
1.10.6 MCL-1 in the Ovary
The pro-survival factor Mcl-1 was found to be expressed in fetal human oocytes of the
primordial follicle pool, oocytes of preantral follicles and granulosa and theca cells of growing
follicles [192, 193]. Upon gonadotropin stimulation there was an increase in follicle survival,
concurrent with an increase in Mcl-1 expression, which suggests that this Bcl-2 family member
may play a role in tilting the balance of follicle fate towards survival [193]. Recently, Jones et al
have confirmed expression of MCL-1 in primordial follicle oocytes and granulosa and via use of
neutralizing antibodies have proposed the necessity for MCL-1 in primordial follicle survival
[181].
46
Until now, a verifiable Mcl-1-deficient oocyte phenotype analysis has not been conducted.
Although the anti-apoptotic function of Mcl-1 in ovaries has been hinted at thus far, a variety of
in vitro and in vivo studies mentioned above strongly suggest additional metabolic and
autophagic roles for Mcl-1 in oocyte and follicle cell fate. We thus propose to establish the
various contributions of Mcl-1 in mediation of oocyte survival and folliculogenesis.
1.11 THESIS HYPOTHESIS AND OBJECTIVES
Oocyte quality and oocyte loss are two factors that determine the reproductive capacity and
length of fertility. The majority of oocytes are lost to PCD and do not survive until ovulation.
Additionally, components of the intrinsic pathway of PCD, the majority of pro-survival factors
of the Bcl-2 family, have been demonstrated to have limited or no impact on postnatal oocyte
survival. MCL-1 expression was localized to fetal human oocytes and the oocytes of primordial
and growing follicles, and an increase in Mcl-1 transcript expression was associated with
increased follicle survival upon gonadotropin stimulation [192, 193].
My overall hypothesis was that Mcl-1 is involved in the regulation of oocyte survival, and the
maintenance of the ovarian reserve, and is modulated by known signaling pathways associated
with follicle survival, placing it as the key survival factor mediating oocyte and follicular fate.
My overall objective was to identify the involvement of Mcl-1 in oocyte survival by
characterization of the Mcl-1 conditional oocyte-specific knockout. Additionally, I wanted to
identify upstream regulators of MCL-1 function in oocytes, and the downstream impact of Mcl-
1-ablation on apoptosis, autophagy and mitochondrial function.
47
As part of the first objective, presented in Chapter 2, I characterized the expression of MCL-1
in oocytes of the neonatal and post-pubertal mouse ovary, and the impact of oocyte-specific
deletion of Mcl-1 on primordial follicle and growing follicle survival, using histomorphometric
analyses, and markers of autophagy, apoptosis and mitochondrial function. In the second
objective, presented in Chapter 3, I looked at granulosa cell-directed mechanisms associated
with mediation of oocyte survival, and their resulting control of MCL-1 expression. Furthermore,
I looked at the putative MCL-1 regulation of mitochondrial metabolic output, proposed by
Perciavalle et al., [221] via the newly defined mitochondrial matrix-restricted MCL-1Matrix
isoform.
48
2 ASSESSING THE ROLE OF ANTI-APOPTOTIC BCL-2
MEMBER MCL-1 IN OOCYTE AND FOLLICLE FATE
2.1 INTRODUCTION
Understanding factors that mediate oocyte and embryo survival are essential in order to contend
with issues of oocyte quality or premature oocyte depletion. The majority of germ cells that
comprise the ovary at birth do not survive to ovulate. In fact, estimates show that 99.9% of germ
cells are eliminated via activation of programmed cell death (PCD) [1, 2]. Insufficient
endowment or excessive oocyte loss, can lead to a premature exhaustion of the ovarian follicle
pool [3]. This leads to a condition known as premature ovarian failure (POF), a syndrome that
affects around 1% of all women. Untimely depletion of the ovarian follicle pool can be caused
by either genetic abnormalities (X-linked or autosomal recessive mutations), iatrogenic factors
(chemotherapy, radiation therapy), or external environmental exposures (various chemical
pollutants), that reduce or abrogate the starting number of oocytes, or disrupt normal follicle
dynamics.
Oocyte susceptibility to PCD can occur at various timepoints during naturally occurring
folliculogenesis [1, 11-13, 15-17, 21, 24]. These ‘windows’ of susceptibility include: primordial
germ cell migration to the embryonic gonad; formation of individualized primordial follicles
with collaborative somatic (granulosa) cell infiltration; entirety of growing follicle pool prior to
post-pubertal rescue by FSH; and selection from antral follicle pool for final ovulation. Whereas
fetal oocytes have long been ascribed to undergo apoptotic cell death, follicular death occurs via
a process termed follicular atresia, characterized by granulosa cell death and recession resulting
in starvation of the oocyte [143, 146, 147]. In vivo assessments of growing follicle pools have
49
revealed that follicular atresia occurs with combined expression of markers of both apoptosis and
autophagy [142]. Once ovulated, oocytes are able to maintain metaphase II arrest for
approximately 24hrs and then begin to develop an increased tendency for spontaneous activation
and cellular fragmentation, classified as containing hallmarks of apoptotic cell death [148, 259].
Additional studies have verified that oocyte-granulosa cell contact is essential for oocyte
metabolic support and the maintenance of meiotic arrest [77-79]. The resumption of meiosis is
one of the first steps preceding oocyte atresia [260], and inhibition of meiotic resumption, even
during DNA damage-inducing death stimuli, prevents the activation of apoptotic cellular
fragmentation [261].
Oocyte quality is an additional factor that can lead to impaired follicular development and
reduced fertility. Maternal age is one particular aspect that contributes to the quality of the
oocyte and the resulting embryo [4]. Older oocytes and zygotes have been well documented to
have increased rates of aneuploidies, oxidative damage, mitochondrial and chromosomal
abnormalities and increased fragmentation rates [4-7]. Aged oocytes have also been associated
with decreased DNA repair [262] as well as impaired mitochondrial function accompanied by
decreases in reactive oxygen species (ROS) and ATP production [119, 120]. Mitochondria have
long been considered the ‘powerhouses’ of the cell, supplying the cell, in this case the oocyte,
with an adequate source of ATP for survival. Mitochondrial number are believed to increase with
oocyte growth, beginning with a few hundred in primordial germ cells, and increasing to a few
hundred thousand by ovulation [107, 108]. Early studies have suggested that mitochondrial
content, specifically mitochondrial DNA (mtDNA) copy number can influence oocyte
developmental competence and fertilization [112, 113]. High levels of ROS have been
implicated in the initiation of various deleterious effects due to oxidative damage, especially on
mtDNA [106], resulting in a reduction in ATP and subsequent meiotic spindle defects in oocytes
50
[114-118]. Thus, fully understanding the mechanics that govern features of proper oocyte
development, specifically with regards to oocyte quality and prevention of premature follicular
depletion, requires us to understand factors that regulate germ cell integrity and survival.
One of the most well studied families of factors responsible for PCD are the anti- and pro-
apoptotic members of the Bcl-2 family. Anti-apoptotic Bcl-2 family members have been
associated with the binding and resultant inhibition of Beclin-induced autophagy (PCD type 2)
both in vivo and in vitro; whereas binding to pro-apoptotic members was not observed [204, 205,
218, 219]. With regards to PCD type 1, or apoptosis, core pro-apoptotic Bcl-2 members are able
to oligomerize, bind and form channels in the mitochondrial membrane triggering the apoptotic
cascade, whereas the anti-apoptotic Bcl-2 members neutralize their killing potential via direct
interaction [126-128]. Additional pro-apoptotic members of the Bcl-2 family can be structurally
variable, and either bind and activate core pro-apoptotic members or inhibit the action of the pro-
survival members [127, 128, 130]. A number of these factors have been identified with specific
functions in the ovary [124].
Disruption of pro-apoptotic Bax in the ovary has been well characterized leading to increased
germ cell survival and prolongation of fertility [152-155, 161]. The involvement of anti-
apoptotic factors have been less clearly defined, with the ablation of Bcl-2, Bcl-w and Diva
(Bcl2l10), having virtually no ovarian phenotype [179-181, 190, 191]. Conditional Bcl-x
inactivation results in a reduced number of fetal germ cells by contributing to an increase in germ
cell apoptosis during embryonic development (somewhere between 12.5dpc and 15.5dpc in the
mouse [158]), however postnatal inactivation of Bcl-x in oocytes did not compromise the ovarian
reserve in young females [187]. Consequently, no Bcl-2 family has been associated with a role in
governing post-natal oocyte survival and maintenance of the adult ovarian reserve.
51
Here we show that an oocyte-specific deletion of Mcl-1 results in a sharp decline in the number
of surviving primordial follicles shortly after birth, and an almost complete depletion of the
ovarian reserve by 3 months of age. Those oocytes that do survive until the early antral follicle
stage display increased mitochondrial malfunction, in addition to elevated hallmarks of
autophagy. Histomorphometric analyses reveal that depletion of the follicular pool in Mcl-1
oocyte-deficient mice can be rescued with concomitant ablation of pro-apoptotic Bax. Thus we
show that Mcl-1 is essential for oocyte survival at all stages, with roles in primordial follicle
survival, maintenance of mitochondrial function and inhibition of oocyte autophagy and post-
ovulatory oocyte death.
2.2 MATERIALS AND METHODS
2.2.1 Animals
For oocyte-specific deletion of Mcl-1, C57BL/6 Mcl-1tm3Sjk
(Mcl-1f) [226] mice carrying the
floxed allele were kindly gifted from the breeding colony of Dr. Joseph T. Opferman and were
intercrossed to mice carrying the Tg(ZP3Cre)3Mrt (Zp3-Cre) transgene [263] (backcrossed for 5
generations from ICR/129 onto C57BL/6 background) resulting in the generation of an oocyte-
specific Mcl-1t2Sjk
(Mcl-1null
) allele. To assess timing of excision, ZP3-Cre mice were crossed to
mice carrying lacZ/alkaline phosphatase reporter line Tg(CAG-Bgeo/ALPP)1Lbe (Z/AP) [264]
and fluorescent reporter line Gt(ROSA)26Sortm9(CAG-tdTomato)Hze
(tdTomato) [265]. For assessment
of oocyte-specific Mcl-1 deletion, female mice of a Mcl-1f/null
: Zp3-Cre allelic compositions
(hereafter referred to Mcl-1cKO) were utilized in our studies to ensure complete early excision
rather than use of Mcl-1f/f
: Zp3-Cre mice. Additionally, Mcl-1f/null
, Mcl-1f/f
, Mcl-1+/+
, Mcl-1+/+
:
Zp3-Cre and Mcl-1f/+
: Zp3-Cre females were collected as controls. Baxtm1Sjk
(Bax-) [150] and
52
Bcl2l11tm1Sjk
(Bim-) [266] knockout animals were obtained from the breeding colonies of Dr.
Jonathan Tilly and Dr. Razq Hakem. Animals were genotyped for possession of either Mcl-1+ or
Mcl-1f alleles using primers CTGAGAGTTGTACCGGACAA (7MCL1) and
GCAGTACAGGTTCAAGCCGATG (6MCL1), and for Mcl-1null
allele using primers 7MCL1
and ACGCTCTTTAAGTGTTTGGCC (2MCL1). Presence of Zp3-Cre transgene was assessed
using TGATGAGGTTCGCAAGAACC (CREF) and CCATGAGTGAACGAACCTGG (CRER)
and genotyping for Bax+ and Bax
- alleles utilized GAGCTGATCAGAACCATCATG (BAX-
EX5-F), GTTGACCAGAGTGGCGTAGG (BAX-LN5-R) and CCGCTTCCATTGCTCAGCGG
(BAX-NEO). Genotyping for Bim+ and Bim
- alleles used CATTCTCGTAAGTCCGAGTCT
(BIM-PB20-COM), GTGCTAACTGAAACCAGATTAG (BIM-PB335-WT) and
CTCAGTCCATTCATCAACAG (BIM-PB65-TA). All mice were housed with free access to
food and water and maintained on a 12h:12h light-dark cycle. All mouse experiments were
performed in accordance with the Canadian Council on Animal Care (CCAC) guidelines for Use
of Animals in Research and Laboratory Animal Care, under protocols approved by animal care
committees at Mount Sinai Hospital (MSH) or the Toronto Centre for Phenogenomics (TCP).
For breeding performance, dams were setup with wildtype male studs with proven breeding
efficiency, and checked for signs of pregnancy and delivery for all litters for a period of 6
months.
2.2.2 Collection of MII and GV Oocytes
Mice were primed by stimulation with 10U PMSG (Pregnant Mare Serum Gonadotropin; NHPP,
USA or ProSpec, Israel (HOR-272)) and 10U hCG (Human Chorionic Gonadotropin; Sigma) 44-
48hrs later. Mature ovulated oocytes were collected using glass pipettes, in mHTF (modified
53
Human Tubal Fluid; Life global) from the oviducts 14-16hrs after hCG. Cumulus cells were
stripped by short (~1 min) incubation in 0.03% Hyaluronidase (Sigma) and then washed in
mHTF.
PMSG-primed ovaries from mice 40-48hrs after stimulation were collected in mHTF and pierced
with a small gauge needle releasing antral cumulus-oocyte-complexes (COC). Denuded oocytes
were collected by manually stripping cumulus cells from these COC’s using narrow bore glass
pipettes. Diploid oocytes arrested in diplotene stage of prophase I are characterized by a large
nucleus termed the germinal vesicle (GV). After ovulation, the haploid oocyte arrests in
metaphase II (MII) in preparation for fertilization.
2.2.3 Histological Analyses
Ovaries from Mcl-1f/null
: Zp3-Cre, Mcl-1f/null
, Mcl-1f/f
, Mcl-1+/+
, Mcl-1+/+
: Zp3-Cre, Mcl-1f/+
:
Zp3-Cre, Bax-/-
, Mcl-1f/+
: Bax-/-
: Zp3-Cre, Mcl-1f/null
: Bax-/-
: Zp3-Cre (hereafter known as Mcl-
1c/Bax DKO) and Mcl-1f/null
: Bim-/-
: Zp3-Cre (Mcl-1c/Bim DKO) females were collected at
varying timepoints (Post natal day 180 (PN180/6 months), PN90 (3 months), PN21 (3 weeks),
PN14 (2 weeks) or PN7) and fixed in Dietrichs (4% Formalin, 28% EtOH, 0.34N Glacial Acetic
Acid (Sigma)) or 10% Formalin (Fisher) and following standard dehydration protocols were
embedded in paraffin wax and sectioned (5µm) using a LEICA RM2255 Microtome and then
mounted on Superfrost plus (Fisherbrand) slides. Sections fixed in Dietrichs were rehydrated and
stained with a picric acid/methyl blue stain, allowing for better resolution for histomorphometric
analyses. Every third section was counted for PN7 and PN14 ovaries, every fifth section counted
for PN21 ovaries, and every 10th
section for PN90 and PN180. Oocytes with visible nuclei from
54
primordial, primary, secondary and antral follicles were quantitated and recorded and multiplied
by associated factor (x3, x5, x10) to gain an approximately full representation of the ovary. For
calculation of rates of atresia, atretic follicles of both advanced stage atresia, and mild to
moderate atresia were quantitated in sections from PN21 ovaries. Follicles in which granulosa
cells displayed the beginnings of death and withdrawal were characterized as mildly atretic, with
moderately atretic follicles displaying a more exacerbated withdrawal. Those follicles in which
the oocyte had completely shrunken were characterized as late stage atretic follicles. To
accurately depict rates of atresia, atretic follicle numbers were taken as a proportion of the total
post-secondary growing follicle pool in each section, per genotype.
Ovaries from 17.5dpc and PN3 tdTomato : Zp3-Cre animals were removed from animals and
washed in mHTF. Entire tissue samples were viewed under LEICA DMI60003 Spinning Disc
Confocal or LEICA MZ 165A Stereomicroscope using TRITC-Red laser (561 excitation, 620
emission), and imaged. Ovaries from Z/AP: Zp3-Cre mice were first fixed 4hrs in LacZ fixative
(0.2% gluteraldehyde, 5mM EGTA pH7.3, 2mM MgCl2, in PBS) on ice and then rinsed in PBS.
Endogenous alkaline phosphatase (AP) activity was inactivated by heating for 30 min in PBS at
70ºC and then rinsed in PBS at RT. Ovaries were then washed in AP buffer (100mM Tris-HCL
pH 9.5, 100mM NaCl, 10mM MgCl2) for 10 min. Ovaries were stained with BM Purple AP
substrate (Roche) at 4ºC for 0.5-36hrs and then washed in 0.1%Tween20 and 2mM MgCl2 in
PBS and sectioned.
Sections fixed in 10% formalin were rehydrated and used for immuno-histochemical staining
protocols. Sections were submitted to antigen retrieval at ~95ºC for 10 min in sodium citrate
buffer (10mM tri-sodium citrate (Sigma) pH 6.0 with HCl), washed and blocked in 10% Normal
Horse Serum (NHS) for 1hr before overnight incubation in primary antibody (in 10%NHS) at
55
4ºC. Primary antibodies utilized include Mcl-1 (Rockland Immunochemicals, 600-401-394S).
Sections were then washed in PBS and incubated with secondary protocols from the ABC
Vectastain kit (PK-4001; Vector Labs) and then visualized using diamino-benzidine (DAB)
(Sigma) substrate. After time-sensitive stain development, sections were counter-stained in
hematoxylin (Sigma) for identification of cell nuclei.
2.2.4 TUNEL Assays
Sections of PN day 1 ovaries from Mcl-1f/null
: Zp3-Cre, Mcl-1f/null
and Mcl-1f/+
: Zp3-Cre females
fixed in formalin were rehydrated as previously described. These sections were then incubated in
15ug/ml Proteinase K in 1X PBS for 15 min and then washed 6 times in 1X PBS. Sections were
then incubated in 0.1% Triton-X in 1X PBS for 10 min and washed again in 1X PBS. Positive
control sections were attained by incubating slides in DNAse1 mix (0.02U/ul DNAse (Sigma),
1X REAct1 buffer (Invitrogen)) for 10 min at RT and then washed in PBS. Slides were
incubated in TdT (Terminal Transferase) Reaction Mix (4uM biotin16-dUTP (Roche), 1.5uM
dATP, 1X NEB4 buffer, 4U/ul TdT enzyme (Roche)) for 1.5hrs in prewarmed container at 37ºC.
Negative control was incubated in Reaction mix without TdT enzyme. Slides were washed in
PBS, and then incubated according to secondary protocols from ABC Vectastain (PK-4001;
Vector Labs) and visualized with DAB, as previously described.
2.2.5 Ovulation Rates, Fragmentation Rates and Breeding Performance
Mice of all genotypes at PN180, PN90 or PN21 were primed and mature ovulated MII oocytes
were collected 14-16hrs after hCG, as described previously, and ovulation rates per female were
56
recorded. For determination of oocyte susceptibility to death when cultured in vitro, MII oocyte
fragmentation rates were recorded. MII oocytes of all genotypes were collected from primed
females at PN21-28, and cultured in HTF (Human Tubal Fluid, Life Global) for a period of
24hrs. The total number of MII oocytes that underwent cellular fragmentation was recorded. For
breeding performance, Mcl-1cKO dams and controls at PN35-42 were mated to young wildtype
males until 6 months of age. Litter sizes and total number of litters were recorded over that
period.
2.2.6 Mitochondrial Analyses – Live cell stains
MII ovulated oocytes of Mcl-1cKO and controls were collected and subjected to a number of
assays for determination of mitochondrial function. Total and respiring mitochondria were
stained using Mitotracker fluorescent dyes (Mitotracker Green FM (M7154), Mitotracker Red
580 (M22425); Molecular Probes, Invitrogen) added to HTF in 100nM concentration for 30min,
following Mitotracker protocols provided, then washed in mHTF and imaged. For total cellular
levels of reactive oxygen species (ROS), oocytes were incubated in 10µM 2’, 7’-
dichlorofluorescein diacetate (DCFDA) dye (DCFDA Cellular Reactive Oxygen Species
Detection Assay Kit (ab113851), Abcam) in HTF for 15 min as per protocol instructions, washed
in mHTF and imaged. Mitochondrial derived superoxides were measured utilizing the MitoSOX
Red fluorescent dye (MitoSOX Red (M36008); Molecular Probes, Invitrogen). MII oocytes from
Mcl-1cKO and controls were incubated in 5µM MitoSOX in HTF for 10 min as in protocol
provided, then washed and imaged. The efficacy of dyes utilized for detection of ROS, or
mitochondrial superoxides was validated previously in our lab, using oocytes cultured in the
presence of various inhibitors of the electron transport chain. Autofluorescence of mitochondrial
57
proteins (NADH and flavoproteins) has been previously utilized to measure the redox state of the
mitochondria [87, 267-269]. Live MII oocytes were imaged for emitted blue autofluorescence of
reduced NADH and NAD(P)H and emitted green autofluorescence indicative of oxidized
flavoproteins (FAD2+). For identification of lysosomes, MII oocytes were incubated with 50nM
Lysotracker Red (LysoTracker Red DND-99 (L7528), Molecular Probes, Invitrogen) in HTF for
30 minutes, as indicated in protocol provided, then washed and imaged. Live oocytes from all
above assays were imaged in mHTF medium.
2.2.7 Immunofluorescence Staining
GV or MII oocytes from Mcl-1cKO and control females were fixed in 10% formalin for 10
minutes and used for staining of markers of apoptosis and autophagy. Oocytes were first
transferred to cooling ~95ºC sodium citrate buffer for antigen retrieval for 10 min using pulled-
glass pipettes. Oocytes were moved to three washes in 0.1% Triton-X in 10mM PBS (0.1%TX)
and then blocked in 10%NHS in 10mM PBS for 15 min. Following this step, oocytes were
incubated in primary antibody in 10%NHS in 10mM PBS overnight at 4ºC. Primary antibodies
used include anti-Beclin-1 (Santa Cruz sc-11427), anti-LC3 (MBL PM046), anti-Lamp1 (1D4B,
Developmental Studies Hybridoma Bank), anti-Lamp2 (ABL-93, Developmental Studies
Hybridoma Bank), anti-Bax-NT (Upstate 06-49), anti-Tubulin (Invitrogen A11126), anti-
phopho-H2AX (Cell Signaling 9718S), anti-Actin (Santa Cruz, sc-1616), anti-AIF (Santa Cruz,
sc-9416), and anti-Mcl-1 (Rockland Immunochemicals 600-401-394S). After primary antibody
incubation, oocytes were transferred to 0.1%TX washes and incubated in secondary antibody in
2% NHS in PBS for 30 min. Secondary antibodies used include host-specific Alexa Fluor dyes
(Invitrogen). Oocytes were washed in 0.1%TX, and shifted to droplet with blue fluorescent 4',6-
58
diamidino-2-phenylindole (DAPI, Sigma) for nuclear stain for 10-15 min. Oocytes were then
mounted on Superfrost slides in 50% glycerol for imaging.
For evaluation of apoptotic induction, Cytochrome c release, AIF release and pan-caspase
activity assays were performed on GV and MII oocytes with protocols modified from BIOMOL
Carboxyfluorescein Multi-Caspase Activity Kit and Cytochrome c release kit (InnoCyte-
Calbiochem). A portion of denuded GV and MII oocytes were permeabilized for 10 min with
digitonin buffer provided in the Cytochrome c release kit. Both permeabilized and non-
permeabilized oocytes were then fixed in fixative supplied, and processed through the staining
protocol provided. For AIF release, AIF (Santa Cruz, sc-9416) was utilized instead of
Cytochrome c antibody provided with kit. Permeabilized and non-permeabilized oocytes were
then incubated with DAPI for 10-15 min and mounted on Superfrost slides in 50% glycerol for
imaging. Total AIF/Cytochrome c released by mitochondria was assessed by comparison of non-
permeabilized AIF/Cytochrome c fluorescence intensity indicating total cellular
AIF/Cytochrome c, to permeabilized AIF/Cytochrome c fluorescence intensity, thereby revealing
the fraction of AIF/Cytochrome c retained in the mitochondria once cell wall was permeabilized.
For pan-caspase activity, GV and MII oocytes were incubated in FML-VAD-FMK stock
dissolved in HTF medium for 2.5hrs and washed and transferred to fixative as indicated in
Caspase activity kit. Oocytes were then washed and transferred to DAPI for 10-15 min, then
mounted on Superfrost slides in 50% glycerol and imaged, as mentioned previously.
59
2.2.8 Imaging
Live or mounted and stained oocytes were imaged using LEICA DMI60003 Spinning Disc
Confocal microscope. Lasers for the following wavelengths were used: DAPI-blue (405
excitation, 450 emission), FITC-green (491 excitation, 525 emission), TRITC-red (561
excitation, 620 emission). Images were acquired and analyzed using Volocity software
(PerkinElmer) with Z-stack images taken at 0.354µm increments across 10µm sections to either
side of the midpoint of the oocyte. Image quantitation was also performed using Volocity
software with consistent quantitation parameters maintained within experiments. Images were
deconvolved using Huygens Essential Software. Images utilized for colocalization data were
analyzed using Imaris (Bitplane) software which additionally supplied us with colocalization
statistical measures.
2.2.9 Metabolic Profile
To assess metabolic profile and determine ATP, citrate, malate and fumarate levels, MII oocytes
were first frozen on glass slides by dipping in cold isopentane which was equilibrated in liquid
nitrogen. Oocytes were then freeze-dried and sent to the lab of Dr. Kelle Moley for processing of
metabolic profile using the protocol delineated in Chi et al. [270].
2.2.10 Statistics
Data were analyzed using either one-way ANOVA with the Holm-Sidak multiple comparisons
test (breeding performance, histomorphometric analyses (PN90)); or using the conservative non-
parametric Kruskal Wallis one-way ANOVA on ranks, followed by Dunns post-hoc test for
60
comparisons between groups, when normality failed or sample sizes were vastly different
(ovulation rates, active Bax (GV), Beclin foci, LC3, MitoTracker Green and Red γH2AX, active
Bax (MII), fragmentation rates). Statistical measures on two samples were performed using the
unpaired t-test (histomorphometric analyses (PN21, PN7), atresia rates, metabolites, MitoSox,
ROS, Cytochrome c, caspase activity).
2.3 RESULTS
The majority of germ cells are destined to undergo PCD [1, 2], and only a few Bcl-2 family
members have been identified with roles in regulation of germ cell fate [158, 179-181, 187, 190,
191], with none attributed to post-natal oocyte survival. As Mcl-1 expression was localized to
fetal germ cells, oocytes of primordial and growing follicles, and granulosa cells of growing
follicles [192, 193], in addition to strong gene expression in ovulated human and murine oocytes
[121], it became apparent that Mcl-1 could be considered a strong candidate for this role.
We have verified expression of MCL-1 in primordial follicle oocytes of PN4 ovaries, and in the
primordial and growing oocyte pool by 3 weeks (PN21) (Fig. 2.1A). Immunoreactivity of MCL-
1 in oocytes grows stronger with the activation of follicle growth (from the primordial to primary
transition), remains robust in the fully grown oocytes of pre-antral and antral follicles and
virtually disappears from oocytes undergoing atresia (Fig. 2.1A). This pattern strongly suggests
MCL-1 as either a regulator of oocyte survival, or a marker of oocyte growth. In order to
establish the functional need for Mcl-1 we created a mouse model with oocyte-specific excision
of Mcl-1 using the Zp3-Cre transgene. We confirmed spatial and temporal expression of the Cre
transgene by intercrossing with two tdTomato and Z/AP reporter lines (Fig. 2.1B). TdTomato:
61
Zp3-Cre ovaries displayed the activation of Cre excision in oocytes as early as 17.5dpc, with a
large proportion of primordial follicle oocytes expressing tdTomato reporter by PN3. The
selectivity of excision was also confirmed in the Z/AP reporter: Zp3-Cre ovaries, in which
excision occurred in a majority of primordial follicle oocytes and virtually all growing oocytes at
3 months of age (PN90). Effective excision of Mcl-1 by the Zp3-Cre transgene was confirmed by
immuno-histochemical (IHC) staining and western blots (WB) (Fig. 2.1C). IHC stains of 3 week
ovaries with MCL-1 antibody revealed strong expression in oocytes within growing follicles of
wildtype ovaries and a drastic reduction in MCL-1 intensity in ovaries from the Mcl-1cKO. This
was supported by the reduction of MCL-1 observed in WBs performed on isolated antral GV
oocytes collected from 3-4 week Mcl-1cKO ovaries.
2.3.1 Breeding Performance, Ovulation Rates and Histomorphometric Analyses
To determine the impact of Mcl-1 oocyte-specific deletion on fertility, Mcl-1cKO and control
females were bred to wildtype control males (proven breeders) starting at 5-6 weeks of age for a
period of 6 months. Mcl-1cKO females had an average of 2 litters, with average litter size of
2.5±0.5 pups as compared to controls that had an average of 5 litters, with average litter size of
6-7 pups. The cumulative breeding performance of Mcl-1cKO dams was thus dramatically
reduced (Fig. 2.2A). Additionally, Mcl-1cKO females did not deliver any further live litters
beyond 4 months of age, whereas all control females of the various indicated genotypes were
able to breed beyond 1 year of age.
In order to establish whether this reduction in fecundity was due to an overall reduction in the
ovulatory capacity, Mcl-1cKO and control females were primed with external gonadotropins and
62
63
Figure 2.1. Patterns of MCL-1 Expression and Verification of Mcl-1 Oocyte-Specific
Excision.
(A) Staining of MCL-1 in ovarian sections of wildtype females (Image Magnification=1000X). Positive
Immuno-histochemistry stain appears as a reddish brown stain whereas nuclear hematoxylin counterstain
appears bluish purple. (i) Postnatal day 4 (PN4) ovaries with some indicated stained primordial follicle
(PMF) oocytes (arrows) or lacking stain (arrowheads). (ii) Immunostainings of 3 week (PN21) ovaries
with marked PMF oocytes (arrows), and indicated stains in growing oocytes of primary and preantral
follicles (arrowheads) and oocytes beginning to undergo early stages of atresia (red arrowhead). (B)
Confirmation of oocyte-specific Zp3-Cre excision using tdTomato Reporter and Z/AP Reporter line. Cre
excision results in red fluorescent stain in oocytes of Tomato Reporter (white arrows) (Image
Magnification=100X) and a dark purple stain in Z/AP Reporter (black arrows) (Image Magnification=400X).
(i) Ovaries from embryonic day 17 (E17dpc) females of the Tomato Reporter line containing Trasgenic
Cre (left) and without Cre (right) and respective brightfield images below. (ii) Ovaries from PN3 Tomato
Reporter line with Cre (left) and without Cre (right) with respective brighfield images below. (iii) Z/AP
Reporter line ovaries from 3 week (PN21) females counterstained with nuclear hematoxylin which
appears pinkish purple. (C) MCL-1 expression in Mcl-1f/-
:Zp3-Cre (Mcl-1cKO) oocytes and controls. (i)
Immuno-histochemistry stain of PN21 Mcl-1cKO (left) and Mcl-1+/+
:Zp3-Cre ovary (right) (Image
Magnification=200X). Intensity of stain reveals relative MCL-1 expression in growing oocyte pool of Mcl-
1cKO (arrowheads) and control ovaries (arrows); and atretic follicles of wildtype controls (red
arrowhead). These results represent replicate immuno-stainings from at least 3 ovaries (ii) Western Blots
(WB) of 200 isolated growing follicle GV oocytes. Membranes incubated with anti- MCL-1 in Mcl-1cKO
oocytes (left) and wildtype control (right), and with anti-ACTIN used as internal control.
64
their ovulatory response quantitated. We used females of various age groups (at 6 month, 3
month and 3 week) as these represented females of middling reproductive age, young
reproductive age and at pubertal onset, respectively. At 6 months, Mcl-1cKO females ovulated
extremely poorly with average ovulation rate of 0.25±0.25 oocytes as compared to an average of
20-23 oocytes in age-matched control genotypes (Fig. 2.2B). Histological observations of
ovarian sections revealed an apparent absence of follicles of all stages of growth and a drastic
reduction in ovary size. The diminished ovulatory capacity was also apparent as early as 3
months, where Mcl-1cKO females ovulated an average of 1.95±0.86 oocytes compared to 25-30
oocytes ovulated by control females (Fig. 2.2B). This reduction translated to a dramatically
impaired ovarian reserve, where Mcl-1cKO females displayed a severe depletion in the number
of primordial follicles, and a limited pool of growing follicles (Fig. 2.2C). Interestingly, at the
onset of puberty (3 weeks), Mcl-1cKO and control females of all genotypes ovulated comparable
numbers of oocytes (Fig. 2.2B); however histomorphometric analyses revealed a sharp reduction
in primordial follicles and a significant decrease in primary follicles (Fig. 2.3A). Intriguingly,
Mcl-1cKO females at 3 weeks displayed no change in atretic follicle proportion of the total
growing follicle pool indicating that the lack of Mcl-1 did not result in an increase in follicular
atresia rates (Fig. 2.3A). However, of those atretic follicles, Mcl-1cKO ovaries do display a
significant increase in the number of late stage atretic follicles which signifies an increasingly
rapid escalation of atresia upon activation of cell death.
The reduction of the primordial ovarian reserve was already established as early as PN7, where
histomorphometric analyses displayed a halving in the number of primordial follicles with no
change in the growing follicle pool (Fig. 2.3B). This apparent lack of impact of Mcl-1-deficiency
on the growing pool at PN7 implied that the follicular depletion was due to increased primordial
follicle loss, rather than an increased growing cohort. This was further confirmed when we
65
66
Figure 2.2. Breeding Performance, Ovulation Rates and Histomorphometric Analyses of
Mcl-1cKO Females and Controls.
(A) Breeding Performance of Mcl-1f/null
: Zp3-Cre (Mcl-1cKO) and control females. (i) Cumulative pup
number from Mcl-1cKO (n=6), Mcl-1+/+
(n=6), Mcl-1+/+
: Zp3-Cre (n=4) and Mcl-1f/f
(n=6) females paired
with wildtype males at PN28 for 6 month breeding trial. Each column represents individual females with
each varied shaded segment indicative of individual litters. (ii) Average litter size of Mcl-1cKO (n=7)
females compared to Mcl-1+/+
(n=13), Mcl-1+/+
: Zp3-Cre (n=6) and Mcl-1f/f
(n=13) control females.
Values represent average number of pups per litter ± SEM. (B) Ovulation rates of Mcl-1cKO and control
females. (i) The number of ovulated oocytes from stimulated 6 month (PN180) Mcl-1cKO (n=12),
compared to Mcl-1+/+
(n=18), Mcl-1+/+
: Zp3-Cre (n=12), Mcl-1f/f
(n=7), and Mcl-1f/null
(n=8) control
females. (ii) Number of ovulated oocytes from stimulated 3 month (PN90) Mcl-1cKO (n=20), compared
to Mcl-1+/+
(n=12), Mcl-1+/+
: Zp3-Cre (n=14), Mcl-1f/f
(n=9), Mcl-1f/null
(n=12), and Mcl-1f/+
: Zp3-Cre
(n=16) control females. (iii) Number of ovulated oocytes from stimulated 3 week (PN21) Mcl-1cKO
(n=21), compared to Mcl-1+/+
(n=19) females. In all age groups, values represent average number of
oocytes ovulated by stimulated females ± SEM. (C) Histological analysis of PN90 Mcl-1cKO compared
to controls. (i) Comparison of largest diameter histological section of Mcl-1cKO and Mcl-1+/+
: Zp3-Cre
ovaries of PN90 females. Ovarian sections stained with nuclear hematoxylin (ii) Histomorphometric
analyses of primordial, primary, secondary and preantral follicles in PN90 Mcl-1cKO (n=5), compared to
Mcl-1+/+
(n=3), Mcl-1+/+
: Zp3-Cre (n=3), Mcl-1f/f
(n=5), Mcl-1f/null
(n=4), and Mcl-1f/+
: Zp3-Cre (n=3)
control females. Values represent average number of follicles per ovary ± SEM. (*= p<0.05, **= p<0.01,
***= p<0.001).
67
Figure 2.3. Histomorphometric Analyses of Mcl-1cKO Females and Controls.
(A) Analyses of 3 week (PN21) ovaries of Mcl-1f/null
: Zp3-Cre (Mcl-1cKO) and control females. (i)
Histomorphometric comparison of primordial, primary and secondary follicle number of Mcl-1cKO (n=5)
to Mcl-1+/+
(n=4) females. Values represent average number of follicles/ovary ± SEM. (ii) Representative
stages of follicle atresia (left) in ovarian sections stained with methyl green, display granulosa cell
death/withdrawal (red arrows) preceding oocyte shrinkage (red arrowhead). Both early and late stage
follicle atresia rates (right) of Mcl-1cKO (n=4), compared to Mcl-1+/+
(n=5) ovaries. Atretic follicles were
taken as a proportion of the total post-secondary growing follicle pool and separated into advanced stage
(late) or early stage follicle atresia. Values represent average percentage of atretic follicles/total secondary
follicles per section ± SEM. (B) Analysis of primordial follicle number in neonates of Mcl-1cKO and
control females. (i) Histomorphometric comparison of primordial, primary and secondary follicle number
of Day 7 (PN7) Mcl-1cKO (n=4), compared to Mcl-1+/+
(n=4) females. Values represent average number
of follicles/ovary ± SEM. (ii) TUNEL stain of PN1 ovaries of Mcl-1cKO (n=4) compared to Mcl-1f/-
(n=3) and Mcl-1f/+
: Zp3-Cre (n=3) females. Ovarian sections (left) were counterstained with methyl green
that appears light blue; with the TUNEL stain (dark brown) marking apoptotic primordial follicles (red
arrowheads). TUNEL positive primordial follicles were counted as a proportion of the total number of
primordial follicles (right). Values represent average percentage of TUNEL positive primordial
follicles/total primordial follicles per section ± SEM. (*= p<0.05, **= p<0.01, ***= p<0.001).
68
performed TUNEL analysis on Mcl-1cKO and controls at PN1. Mcl-1cKO ovaries displayed a
significant doubling of the number of apoptotic (TUNEL positive) primordial follicle oocytes
compared to controls (Fig. 2.3B).
We thus conclude that Mcl-1 is required for postnatal oocyte survival and the maintenance of the
ovarian reserve; as oocyte ablation of Mcl-1 leads to a drastic reduction in the ovarian reserve.
2.3.2 Markers of Apoptosis in Growing Follicle Pool
As Mcl-1 ablation was able to sharply reduce follicle number, displayed an increased pre-
disposition to activation of cell death, and MCL-1 expression was absent from follicles
undergoing atresia, we wanted to assess the fate of the growing follicle pool by analyzing
markers of PCD. As primordial follicle oocytes of Mcl-1cKO ovaries undergo excessive
activation of cell death with hallmarks of apoptosis, we set to investigate whether grown GV
oocytes also exhibit increases in markers of apoptosis. Oligomerization of pro-apoptotic Bcl-2
members result in the formation of a pore spanning the OMM, allowing release of a number of
factors from the intermembrane space into the cytosol [138]. These factors include Cytochrome c
(CYC1), Second mitochondrial activator of caspases (SMAC)/Direct IAP binding protein with
low PI (DIABLO), High-temperature requirement protein A2 (HtrA2)/Omi, Apoptosis Inducing
Factor (AIF), Endonuclease G and Caspase-Activated DNAse (CAD) [123]. Upon release, these
mitochondrial-sequestered factors trigger the apoptotic cascade by jointly mediating activation of
cysteinyl aspartic acid proteases (caspases) and DNAses, for DNA fragmentation, chromatin
condensation and effective proteolytic destruction of cell components.
69
GV oocytes were stained for activation of pro-apoptotic Bcl-2 members in addition to markers of
the apoptotic cascade. Relative to controls, Mcl-1cKO oocytes exhibited an increase in
fluorescent intensity of BAX-NT, utilizing an antibody recognizing oligomerized/activated BAX
(Fig. 2.4A). Intriguingly, this was not associated with an activation of the apoptotic cascade, as
Mcl-1cKO GV oocytes did not display any significant increase in elements of the apoptotic
cascade. Mitochondrial Cytochrome C or AIF release showed no change in Mcl-1cKO GV’s and
this was accompanied by no apparent increase in pan-caspase activity, when compared to
controls (Fig. 2.4B, C, D). This data effectively demonstrates that despite BAX oligomerization,
there is no subsequent instigation of the apoptotic cascade in Mcl-1cKO GV oocytes. Bax has
been previously shown to be dispensable for growing follicle atresia, as Bax-deficient ovaries do
still undergo comparable antral follicle atresia [161]. However, further work is required to
ascertain the role of Bax in follicle atresia of Mcl-1-deficient oocytes; whether BAX activation
represents a novel non-apoptotic function, or if alternative cell death pathways mediate follicle
atresia.
2.3.3 Markers of Autophagy in Growing Follicle Pool
Previous studies have already observed that follicle atresia exhibits hallmarks of both autophagic
(PCD type 2) and apoptotic (PCD type 1) cell death [142]. Mcl-1 has been previously linked to
roles in both inhibition of autophagy [203, 204, 219] and apoptosis [126-128]. Therefore, after
observing no impact of Mcl-1-depletion on the activation of apoptosis in the antral follicle pool,
we turned to the assessment of markers associated with the activation of autophagy.
Macroautophagy is the manner by which cells are able to effectively degrade selected cellular
components for energy in times of starvation. In addition to this role, autophagy has also been
70
demonstrated to be utilized in cellular recycling, by elimination of malfunctioning organelles, or
for quick energy production in times of stress; thus revealing itself as a cell survival pathway.
Initiation of autophagy results in the formation of an autophagosome surrounding the material
recruited for degradation. The autophagosome fuses with one or more lysosomes forming an
autolysosome, the material within is degraded and the resulting nutrients released [194-196].
Gonadotropin-primed Mcl-1cKO and control ovaries were dissociated for collection of denuded
antral follicle GV oocytes as described earlier. These GV oocytes were stained with a number of
markers of autophagy, and fluorescence intensity or numbers of autophagic vesicles were
quantified. Induction or phosphorylation of Beclin-1/BECN-1 has been shown to initiate
autophagosome formation [197, 199, 271], and Mcl-1 has been demonstrated to bind and inhibit
activation of Beclin-1 induced autophagy [204, 219]. Mcl-1cKO oocytes displayed an increase in
size and number of Beclin-1 foci compared to controls, indicating an increase in Beclin-1-
associated vesicle formation (Fig. 2.5A). Additionally, Microtubule-associated protein 1 light
chain (MAP1LC3A), also known as LC3, has been documented with roles in autophagosome
membrane elongation, autophagosome membrane closure and substrate targeting for autophagic
degradation [196, 207, 209]. Mcl-1cKO GV oocytes also displayed a relative increase in
fluorescence intensity of total LC3 (Fig. 2.4B).
Lysosome-associated membrane proteins 1 and 2 (LAMP-1, LAMP-2) are an essential part of
autophagosome maturation and have been found to be vital for auto-lysosome formation and
lysosomal fusion [211]. When quantified, Mcl-1cKO GV oocytes exhibited no change in number
or volume of LAMP1/LAMP2 positive structures, but showed a significant increase in structures
carrying both Beclin-1 and LAMP-2 positivity (Fig. 2.5B). Colocalization analyses confirmed
this significant increase in Beclin-1 and LAMP-2 co-localized foci in Mcl-1cKO GV oocytes.
71
72
Figure 2.4. Markers of Apoptosis and Autophagy in GV Oocytes.
(A) Immunofluorescent stain for apoptotic markers in GV oocytes from 3 week (PN21) ovaries. Isolated
GV oocytes were stained with anti-Active Bax (green) and counterstained with DAPI (blue) to mark
nuclei (left). Images displayed are representative of n=10 Mcl-1f/null
: Zp3-Cre (Mcl-1cKO) GV oocytes (a)
with brightfield image (c) and secondary only negative control (e); and n=16 Mcl-1+/+
GV’s (b) with
brightfield image (d) and negative control (f). The mean signal intensity within each oocyte was
quantitated (right) for each GV oocyte in addition to Mcl-1f/+
: Zp3-Cre (n=7) GVs. Values are displayed
as average relative fluorescence units (RFU x104) per oocyte ± SEM. (B) For the Cytochrome c release
assay, Mcl-1cKO (n=19), Mcl-1+/+
(n=33) and Mcl-1f/-
(n=5) GV oocytes, and permeabilized Mcl-1cKO
(n=13), Mcl-1+/+
(n=31) and Mcl-1f/-
(n=6) oocytes, were stained with anti-Cytochrome c (green) to
determine the proportion of Cytochrome c retained in mitochondria. Mean signal intensity was
quantitated and values represent the average fold change of RFUs per oocyte ± SEM of retained/total
Cytochrome c, normalized to average Mcl-1+/+
value. Differences were not significant. (C) For the AIF
release assay, Mcl-1cKO (n=11), Mcl-1+/+
(n=12) and Mcl-1f/-
(n=8) GV oocytes, and permeabilized Mcl-
1cKO (n=9), Mcl-1+/+
(n=12) and Mcl-1f/-
(n=6) oocytes, were stained with anti-AIF (green) to determine
the proportion of AIF retained in mitochondria. Mean signal intensity was quantitated and values
represent the average fold change of RFUs per oocyte ± SEM of retained/total AIF, normalized to average
Mcl-1+/+
value. Differences were not significant (D) Mcl-1cKO (n=12) and Mcl-1+/+
(n=7) GV oocytes
were assessed for pan-Caspase activity. Mean intensity was quantitated per oocyte and values indicate
average RFUs per oocyte ± SEM. Differences were not significant. (E) Immunofluorescent stain of total
LC-3 in isolated PN21 GV oocytes. Isolated Mcl-1cKO (n=9) and Mcl-1+/+
(n=7) GV oocytes were
stained for total anti-LC3 (red), marker of preliminary autophagosome formation, counterstained with
DAPI, and representative image displayed (left). Mean signal intensity of each oocyte was quantitated
(right) and values shown represent the average relative fluorescence units (RFU x104) per oocyte ± SEM
(F) Transmission Electron Microscopy (TEM) imaging of Mcl-1cKO and wildtype control GV oocytes.
Presence of electron dense structures (inset) characterized as lysosomes (red arrows) observed in Mcl-
1cKO. (**= p<0.01, ***= p<0.001).
73
The increase in auto-lysosome-like structures was further validated by Transmission Electron
Microscopy (TEM) where an increased association in Mcl-1cKO oocytes was noted between
electron dense structures, reminiscent of lysosomes, and what appear to be autophagic vacuoles
(Fig. 2.4C).
The increase in number of Beclin-1-positive vesicles, elevated LC-3 immunoreactivity, and
increases in Beclin-1 and LAMP-2 co-localized structures, indicates an augmentation of
autophagosome and autolysosome formation, triggered due to Mcl-1-deficiency.
2.3.4 Mitochondrial Functionality in Ovulated Oocyte Pool
Although Mcl-1-deficient GV oocytes activate autophagy, this does not lead to excessive follicle
atresia (Fig. 2.3A). One possible causative factor for autophagic activation is impairment in
mitochondrial bioenergetics leading to increased nutrient deprivation or starvation. As oocyte
quality and reproductive competence have been suggested to rely on mitochondrial content and
effective mitochondrial bioenergetic output [112-115], we assessed those specific parameters in
Mcl-1cKO ovulated oocytes and controls.
Ovulated MII oocytes from Mcl-1cKO and control females were stained for markers of
mitochondrial function. Staining with MitoTracker Red, a dye taken up by mitochondria that are
actively respiring, revealed a significant reduction in the respiring mitochondrial pool in Mcl-
1cKO MII oocytes (Fig. 2.6A). This decrease was significant despite a marked elevation in the
total mitochondrial pool evidenced by increased MitoTracker Green intensity, a dye marking all
mitochondrial organelles (Fig. 2.6A). To assess the impact of this significant reduction in
respiring mitochondrial number we measured overall mitochondrial output by measuring levels
74
75
Figure 2.5. Markers of Autolysosome Formation in GV Oocytes.
(A) Immunofluorescent stain of Beclin and LAMP-2 in isolated GV oocytes from PN21 (3 week)
ovaries. (i) GV oocytes were stained with anti-Beclin-1 (red), marker of autophagosomes, and anti-
LAMP2 (green), marker of lysosomes, and counterstained with nuclear fluorescent stain DAPI. Images
are representative of n=19 Mcl-1f/null
: Zp3-Cre (Mcl-1cKO) GV oocytes (a) accompanied by brightfield
image (c) and secondary only negative control (e); and n=13 Mcl-1+/+
GVs (b) with brightfield image (d)
and negative control (f). (ii) Foci structures noted with anti-Beclin in previous image (white arrow),
indicating autophagosome formation, were calculated and all foci larger than 3µm in volume were
counted from all stained GV oocytes in addition to n=13 Mcl-1f/+
: Zp3-Cre GVs. Values represent the
average number of Beclin-1 foci per oocyte ± SEM (iii) The volume of Beclin-1 foci, was also assessed in
Mcl-1cKO compared to controls and the data shown indicates the average volume of Beclin-1 foci per
oocyte ± SEM. (B) 3D rendering of Beclin-1 and LAMP2 stained GV oocytes. To determine co-
localization of Beclin and LAMP2, indicative of increased autolysosomal formation, 3D rendering was
performed on Mcl-1cKO and Mcl-1+/+
GVs stained for Beclin (a and e respectively), LAMP2 (b and f
respectively), and co-localized pixels (c, d and g,h respectively). Coefficients of co-localization were
calculated using IMARIS software for individual Mcl-1cKO oocytes compared to Mcl-1+/+
oocytes
(Pearsons(PCC) = 0.51±0.05 to 0.17±0.07 respectively; p<0.001)(Manders A(MCC-A) = 0.098±0.017 to
0.058±0.01 respectively; p<0.05)(Manders B(MCC-B) = 0.079±0.014 to 0.032±0.005 respectively;
p<0.05). (*= p<0.05, **= p<0.01, ***= p<0.001).
76
of substrates of the citric acid (TCA) cycle and total cellular ATP levels. Mcl-1cKO MII oocytes
showed no apparent difference in total ATP when compared to control, implying no change in
available energy (Fig. 2.6B). Measures of TCA cycle substrates revealed no significant change in
levels of citrate, however Mcl-1cKO oocytes presented a sharp reduction in levels of fumarate
and malate, downstream elements of the TCA cycle (Fig. 2.6B). Thus although Mcl-1cKO MII
oocytes display no apparent deficiencies in available energy sources, levels of TCA cycle
constituents are severely impacted.
Although total cellular ATP levels were unaltered, total cellular ROS levels were markedly
increased in Mcl-1cKO MII oocytes (Fig. 2.7A), and this was additionally supported by
significant increases in MitoSox Red, a fluorescent marker of mitochondrial derived ROS (Fig.
2.7A). High ROS levels can be indicative of defective antioxidant machinery or an
inhibition/block in the electron transport chain [102-105]. Additionally, changes in ROS levels
can be indicative of mitochondrial performance [119, 120], with high ROS levels implying
overuse of the mitochondrial bioenergetics machinery.
Oocyte quality has been suggested to be highly reliant on mitochondrial function [112-115].
Elevated ROS levels, reduction in ATP content and improper oxidative metabolism have been
linked with defective development and meiotic spindle defects in oocytes and resultant embryos
[93, 114-118]. Microtubule staining of the meiotic spindle apparatus revealed increased spindle
defects in addition to elevated rates of chromosomal misalignments in Mcl-1cKO oocytes
compared to controls (Fig. 2.7B). High ROS levels have also been associated with increases in
oxidative damage on both nuclear and mitochondrial DNA [106], however Mcl-1cKO MII
oocytes revealed no elevation in nuclear DNA damage, as demonstrated by γ-H2AX, a widely
used marker of double stranded breaks (Fig. 2.7B).
77
Figure 2.6. Markers of Mitochondrial Functionality.
(A) MII oocytes were stained for markers of mitochondrial function. (i) MII oocytes were stained with
MitoTracker Green, which stains the total mitochondrial population of the cell. Displayed left are
representative images from n=41 Mcl-1f/null
: Zp3-Cre (Mcl-1cKO) MII oocytes (a) with brightfield image
below (c), and n=43 Mcl-1+/+
MII oocytes (b) with brightfield (d). The mean signal intensity from these
MII oocytes and n=25 Mcl-1f/+
: Zp3-Cre oocytes were quantitated (right) and the values graphed
represent average fold change ± SEM of quantitated relative fluorescence per oocyte, normalized to
average Mcl-1+/+
oocyte value. (ii) MII oocytes were also stained with MitoTracker Red, an indicator of
actively respiring/functional mitochondria. Representative images (left) of n=68 Mcl-1cKO and n=52
Mcl-1+/+
MII oocytes are displayed. Mean signal intensity from these in addition to n=10 Mcl-1f/+
: Zp3-
Cre oocytes were quantitated (right) with data shown indicative of average fold change ± SEM of
quantitated relative fluorescence per oocyte, normalized to average Mcl-1+/+
oocyte value. (B)
Mitochondrial output and bioenergetics in MII oocytes. Mcl-1cKO (n=15) and Mcl-1+/+
(n=15) oocytes
were assayed for metabolite levels of ATP or citrate in addition to downstream constituents of the TCA
cycle, malate and fumarate. Values are represented by average metabolite level per oocyte wet weight ±
SEM. (*= p<0.05, **= p<0.01, ***= p<0.001).
78
79
Figure 2.7. Markers of Mitochondrial Functionality, DNA Damage and Spindle Assembly.
(A) MII oocytes were measured for reactive oxygen species (ROS) levels. (i) MII oocytes were stained
with dichlorofluorescein diacetate (DCFDA) dye (green) to stain total cellular ROS. Representative
images displayed (left) of n=20 Mcl-1f/null
: Zp3-Cre (Mcl-1cKO) (a), with brightfield image (c), and n=17
Mcl-1+/+
MII oocytes (b), with respective brightfield image (d). The mean signal intensity of each
individual oocyte was quantitated with the graph (right) showing average fold change ± SEM of relative
fluorescent units per oocyte normalized to average Mcl-1+/+
oocyte value. (ii) MII oocytes were stained
with MitoSox (red), a marker of mitochondrial derived superoxides. Images displayed (left) are a
representative image of n=20 Mcl-1cKO and n=24 Mcl-1+/+
oocytes. Mean intensity was quantified from
each oocyte (right) with graph values representing relative fluorescent units (RFUx106) per oocyte ±
SEM. (B) MII oocytes were assessed for DNA damage and disrupted spindle assembly. (i) MII oocytes
were stained with DAPI nuclear stain to visualize chromosomes. Number of chromosomes not aligned on
metaphase plate were counted in Mcl-1cKO (n=27) and Mcl-1+/+
(n=41) MII oocytes. Values represent
percentage of chromosomes misaligned compared to total chromosomes analyzed per oocyte. Examples
of chromosomal misalignments are shown in γ-H2AX and Tubulin stained images marked with red
arrows. (ii) Immunofluorescent stains with anti-Tubulin (green) were performed on MII oocytes to
visualize meiotic spindle apparatus and counterstained with DAPI (blue). Images displayed are
representative spindles observed in Mcl-1cKO and control GV oocytes. (iii) DNA damage was measured
in MII oocytes using immunofluorescent stain with anti-phospho Histone H2AX (γ-H2AX) (green), a
marker of DNA double strand breaks, counterstained with DAPI (blue). Images displayed (left) are a
representative sample of n=28 Mcl-1cKO MII oocytes stained with anti-γ-H2AX (a) and DAPI (c), and
n=8 Mcl-1+/+
MII oocytes stained with anti-γ-H2AX (b) and DAPI (d). Mean signal intensity of these
chromosomal stains were quantitated (right), in addition to MII oocytes of Mcl-1f/-
(n=17) and Mcl-1f/+
:
Zp3-Cre (n=20) females. Values represent average relative fluorescence units (RFUx106) per
chromosomal region ± SEM. (***= p<0.001).
80
In summary, deletion of Mcl-1 impacts mitochondrial functionality and bioenergetics. Mcl-1cKO
oocytes carry an increased total mitochondrial population, but with a small but significant
reduction in actively respiring mitochondria. This impairment does not seem to change total
cellular ATP levels; and yet Mcl-1cKO MII oocytes carry a defect in downstream production of
TCA cycle constituents, have elevated ROS levels and exhibit increased incidences of spindle
abnormalities and chromosomal misalignments.
2.3.5 Viability of Ovulated Oocytes
Despite the normal number of oocytes ovulated by Mcl-1cKO females at 3 weeks, breeding
performance was poor. Thus, we decided to investigate whether the viability of these ovulated
oocytes was compromised. Ovulated MII oocytes were collected from PMSG-primed 3 week
Mcl-1cKO and control females and supporting cumulus cells removed. To reconfirm the
continuing activation of the autophagic pathway in ovulated MII oocytes, we stained Mcl-1cKO
MII oocytes and controls with LysoTracker Red, a fluorescent dye that marks acidic organelles
such as lysosomes. Mcl-1cKO MII oocytes had elevated numbers of clearly marked LysoTracker
positive foci when compared to controls (Fig. 2.8A).
In order to determine whether ovulated Mcl-1cKO MII oocytes were exhibiting an increase in
apoptotic markers relative to controls, we subjected them to similar assays that we had
performed on the growing follicle GV oocytes, mentioned earlier. Additionally, in vitro culture
of MII oocytes revealed an inability of Mcl-1cKO MII oocytes to sustain meiotic arrest and an
increased propensity to undergo cellular fragmentation, indicating compromised oocyte survival.
Cellular fragmentation has been characterized as exhibiting hallmarks of apoptotic cell death
81
[148], and nearly 50% of Mcl-1cKO oocytes cultured in vitro for 24hrs fragmented, as opposed
to 5-10% of control MII oocytes (Fig. 2.8C).
Furthermore, Mcl-1cKO MII oocytes displayed an increase in activated BAX, indicating its
oligomerization/activation (Fig. 2.8B). Although in Mcl-1cKO GV oocytes, activation of BAX
was unaccompanied by increase in activation of the downstream apoptotic cascade relative to
controls, Mcl-1cKO MII oocytes did in fact activate the apoptotic cascade. Mcl-1-deficiency led
to an increase in mitochondrial Cytochrome C release (Fig. 2.8B), in addition to increased pan-
Caspase activity (Fig. 2.8C). Thus, Mcl-1 deficiency in ovulated MII oocytes appears to result in
activation of the apoptotic cascade, leading to an increased proclivity for these compromised
oocytes to fragment. As cellular fragmentation has been revealed to contain hallmarks of
apoptotic cell death [148], and inhibition of meiotic progression prevents activation of apoptotic
cellular fragmentation [261], we can assume that this propensity of Mcl-1cKO MII oocytes to
fragment is accompanied by an inability to sustain meiotic arrest.
2.3.6 Rescue of Mcl-1-Deficient Phenotype by Deletion of Bax
Deletion of the pro-apoptotic Bcl-2 family member Bax has been previously established to
rescue the pre-natal primordial germ cell loss induced by ablation of the anti-apoptotic Bcl-x
[158]. By itself, Bax deficiency has been linked to increased primordial germ cell survival [159]
which has been proposed to lead to an increase in follicular endowment [160]. Bax-deficient
females do sustain their ovarian function to advanced chronological age [156, 157], but apoptosis
of post-meiotic oocytes and primordial follicles was not impacted [160]; and Bax has also been
found to be dispensable for follicular atresia [161].
82
83
Figure 2.8. Markers of Autophagy and Apoptosis in MII Oocytes.
(A) MII oocytes were stained with LysoTracker Red, a marker of lysosome formation and indicator of
induction of autophagy (200X). Lysosomal foci formation (white arrows) indicated in Mcl-1f/null
: Zp3-Cre
(Mcl-1cKO) (n=9) and Mcl-1+/+
(n=14) MII oocytes, with accompanying brightfield images below. (B)
MII oocytes were stained for markers of activation of the apoptotic cascade. (i) Representative images of
MII oocytes stained with anti-Bax NT (green), indicative of active BAX oligomerization, and
counterstained with nuclear DAPI, are displayed (left). Images show Mcl-1cKO (n=12), oocytes stained
with anti-Bax (a), with brightfield image (c), and negative control (e); and Mcl-1+/+
(n=13) oocytes
stained with anti-Bax (b), above brightfield (d) and negative control (f). The mean intensity of these
images was quantified, in addition to n=7 and Mcl-1f/-
MII oocytes and values in graph (right) represent
the average relative fluorescence units (RFUx103) per oocyte ± SEM. (ii) Additional downstream factors
of apoptotic cascade, mitochondrial Cytochrome c release was evaluated using anti-cytochrome c
fluorescent stain of MII oocytes. MII oocytes were subjected to cell membrane permeabilization.
Permeabilized and non-permeabilized cells were stained with anti-Cytochrome c (green) (200X) to
determine the proportion of Cytochrome c retained in mitochondria. Images displayed (left) show anti-
Cytochrome c fluorescently stained representative oocyte from n=13 Mcl-1cKO MII oocytes (a) and
permeabilized Mcl-1cKO oocytes (c); in addition to stained n=30 Mcl-1+/+
MII oocytes (b) and
permeabilized Mcl-1+/+
oocytes (d). Mean intensity of these oocytes were quantitated (right) and graph
values represent the average fold change of relative fluorescence unit quantitation per oocyte ± SEM of
Cytochrome c/retained Cytochrome c, normalized to average Mcl-1+/+
value. (C) Analysis of
susceptibility to apoptosis in MII oocytes. (i) MII oocytes were stained for pan-Caspase activity (green) to
ascertain downstream activation of the apoptotic pathway. Images (left) represent single oocyte from Mcl-
1cKO (n=16) MII oocytes and Mcl-1+/+
(n=13) control oocytes (200X). Mean intensity was quantitated
from these oocytes (right) and graph values indicate average transformed relative fluorescence units
(RFUx10-1
) per oocyte ± SEM. (ii) MII oocytes were incubated for 24hrs in HTF culture to determine
fragmentation susceptibility. Mcl-1cKO (n=6), Mcl-1+/+
(n=9), Mcl-1+/+
: Zp3-Cre (n=3), and Mcl-1f/+
:
Zp3-Cre (n=8) females were super-ovulated and predisposition to fragment in culture was documented
and plotted, with values displayed representing average percentage of fragmented oocytes of total oocyte
pool ± SEM per genotype. (*= p<0.05, **= p<0.01, ***= p<0.001).
84
In order to evaluate whether Bax-deficiency would also rescue the primordial follicle phenotype
in Mcl-1cKO females, in addition to the increased susceptibility to death in growing follicles and
ovulated MII oocytes, we crossed a Bax-deficient line to our Mcl-1cKO mouse line. Ovaries
from 3 month females of Mcl-1cKO, Bax-/-
(hereafter known as BaxKO), Mcl-1f/-
: Bax-/-
: Zp3-
Cre (hereafter known as Mcl-1c/BaxDKO) and previously listed controls were collected for
histomorphometric ovarian analyses. BaxKO ovaries revealed a significantly increased cohort of
primordial follicles, verifying the previously described phenotype of enhanced follicle survival
with Bax-deficiency (Fig. 2.9A). Mcl-1c/BaxDKO ovaries exhibited a histomorphometric
phenotype similar to that of BaxKO ovaries, with elevated numbers of follicles of all stages
compared to Mcl-1cKO ovaries, implying a rescue of the Mcl-1-deficient oocyte phenotype (Fig.
2.9A).
We also measured the ovulatory capacity of gonadotropin-primed Mcl-1cKO, BaxKO, Mcl-
1c/BaxDKO and control females at 3 months to assess whether Mcl-1c/BaxDKO ovaries retained
a growing follicle cohort comparable to controls. Ovulation rates of Mcl-1c/BaxDKO were akin
to controls with Mcl-1c/BaxDKO females averaging 28 MII oocytes compared to 20-25 oocytes
as ovulated by BaxKO and control animals (Fig. 2.9B). Comparatively, Mcl-1cKO females
ovulated an average of 2 MII oocytes at 3 months. Moreover, as denuded Mcl-1cKO MII oocytes
had displayed a predisposition to apoptotic cell death by cellular fragmentation when cultured in
vitro for 24hrs; we subjected ovulated MII oocytes from BaxKO and Mcl-1c/BaxDKO to the
same conditions. BaxKO and Mcl-1c/BaxDKO displayed a definitive resiliency against cellular
fragmentation as a majority of cultured oocytes did not fragment (0.03%), when compared to a
near 50% fragmentation rate of Mcl-1cKO oocytes (Fig. 2.10A).
85
Ablation of Bax is thus able to restore the primordial follicle allotment in females with oocyte-
specific deficiency of Mcl-1, as evidenced by a primordial follicle pool comparable to controls.
Additionally, Bax deletion also reduces the increased susceptibility of Mcl-1cKO MII ovulated
oocytes to fragment in culture. In order to determine whether Bax deficiency was also able to
prevent the mitochondrial dysfunction exhibited by Mcl-1cKO oocytes, we stained Mcl-
1c/BaxDKO, Mcl-1cKO and control oocytes with MitoTracker Red, a marker of actively
respiring mitochondria. Mcl-1c/BaxDKO oocytes maintained a significantly lower population of
respiring mitochondria compared to controls (Fig. 2.10B), indicating compromised
mitochondrial functionality. Furthermore, total cellular ATP output in Mcl-1c/BaxDKO
remained unchanged (data not shown), indicating a lack of impact of this reduction of
functionally active mitochondria on total energy supply.
In addition to rescue by Bax-ablation we assessed whether deletion of the pro-apoptotic BH3-
only activator Bim would rescue the Mcl-1-deficient phenotype. BIM, BID and PUMA, have
been identified as BH3-only activators of BAX in in vitro studies [131-134], and Bim expression
was identified in primordial follicle oocytes and granulosa cells of growing follicles [173-175];
which implicates BIM as a putative activator of BAX for regulation of primordial oocyte fate.
Histomorphometric analyses of 3 month Mcl-1c/BimDKO ovaries, revealed no rescue of
primordial, primary or secondary follicle number (Fig. 2.9A), with follicle numbers reminiscent
of Mcl-1cKO ovaries; however, concomitant deletion of Bim did lead to moderate rescue of Mcl-
1-deficient ovulated oocyte number (Fig. 2.9B). Additionally, the increased fragmentation
susceptibility in Mcl-1cKO MII oocytes cultured for 24hrs also appeared to be alleviated by
concurrent Bim deletion. Therefore, Bim, although apparently not required for primordial follicle
cell fate, may play a role in late stage growing follicle atresia, and ovulated oocyte death.
86
Figure 2.9. Rescue of Mcl-1-Deficient Follicle Loss by Concurrent Bax-Ablation.
(A) Histomorphometric analyses of 3 month (PN90) ovaries. Comparison of primordial, primary and
secondary follicle number in PN90 Mcl-1f/null
: Zp3-Cre (Mcl-1cKO) (n=5), Mcl-1+/+
(n=3), Mcl-1f/+
: Zp3-
Cre (n=3), Mcl-1f/+
: Bax-/-
:Zp3-Cre (n=4), Mcl-1f/null
: Bax-/-
: Zp3-Cre (Mcl-1c/BaxDKO) (n=3), and Mcl-
1f/null
: Bim-/-
: Zp3-Cre (Mcl-1c/BimDKO) (n=2) females. Values represent average follicle number per
genotype ± SEM. (B) Ovulation rates of 3 month (PN90) females. Mcl-1cKO (n=20), Mcl-1+/+
(n=12),
Mcl-1f/+
: Zp3-Cre (n=16), Mcl-1f/+
: Bax-/-
:Zp3-Cre (n=5), Mcl-1c/BaxDKO (n=3), and Mcl-1c/BimDKO
(n=2) females were primed with PMSG and hCG and total ovulated MII oocyte pool was counted. Values
represent average number of ovulated oocytes ± SEM per female. (*= p<0.05, **= p<0.01, ***=
p<0.001).
87
Figure 2.10. Impact of Bax-Ablation on Mcl-1-Deficient Oocyte Mitochondrial Function
and Apoptosis.
(A) Analysis of apoptotic susceptibility of MII oocytes. MII oocytes from stimulated Mcl-1cKO (n=6),
Mcl-1+/+
(n=10), Mcl-1f/+
: Zp3-Cre (n=4), Mcl-1f/+
: Bax-/-
:Zp3-Cre (n=2), Mcl-1c/BaxDKO (n=2) and Mcl-
1c/BimDKO (n=2) females were submitted to 24hr culture in HTF medium to determine fragmentation
predisposition. Values represent the average percentage of fragmented oocytes per female ± SEM over
total cultured oocyte pool. (B) MII oocytes were analyzed for markers of mitochondrial function. Mcl-
1cKO (n=50), Mcl-1+/+
(n=33), Mcl-1f/+
: Bax-/-
:Zp3-Cre (n=7) and Mcl-1f/null
: Bax-/-
: Zp3-Cre (Mcl-
1c/BaxDKO) (n=49) MII oocytes were analyzed for mitochondrial functionality using MitoTracker Red, a
marker of respiring mitochondria (200X). Images displayed (left) represent a single oocyte of each
genotype, and mean intensity of all oocytes were quantitated (right), with graph values representing fold
change of average relative fluorescence unit per oocyte ± SEM normalized to average Mcl-1+/+
value.
(**= p<0.01, ***= p<0.001).
88
2.4 DISCUSSION
The lack of an ovarian phenotype with deletion of anti-apoptotic Bcl-2 members Bcl-2, Bcl-w
and Bcl2l10 (Diva) [179-181, 190, 191], in addition to the absence of a postnatal impact of
conditional Bcl-x ablation [187] has led to the postulation that anti-apoptotic Bcl-2 members are
either uninvolved, or play minor or redundant functions in postnatal oocyte survival. The work in
this study presents Mcl-1 as the first documented pro-survival Bcl-2 member required for
postnatal oocyte survival, and maintenance of the ovarian reserve. Mcl-1 has a well-documented
role in cell survival; however its conditional deletion in oocytes has not only confirmed the
involvement of Mcl-1 in decisions of cell fate, but has also revealed its inhibition of varied
modes of oocyte cell death, albeit in a stage-specific manner.
As demonstrated above, cytoplasmic MCL-1 expression increases from the primordial to primary
transition, and continues to aggregate in an association with sustained follicle growth. The
phenotype of primordial follicle loss in neo-natal Mcl-1cKO ovaries, with no associated increase
in the growing follicle pool, implicates Mcl-1 in the mediation of primordial follicle survival.
Further studies are required to assess whether this loss is due to the lack of the anti-apoptotic
impact of MCL-1, or additional roles MCL-1 may play in either DNA repair or cell-cycle arrest
[272, 273]. Subsequent deletion of Bax in this Mcl-1-deficient oocyte model prevents the
primordial follicle loss, resulting in perseverance of the PMF pool comparable to the Bax-
deficient phenotype. Hence, whatever role Mcl-1 does play in regulation of primordial follicle
survival, does appear to be antagonized by Bax action.
Mcl-1-deficient oocytes that escape the early primordial follicle demise and begin to grow,
exhibit increased markers of cellular autophagy and mitochondrial dysfunction without
89
activation of the apoptotic pathway downstream of BAX activation. It is probable that Mcl-1-
deficient GV oocytes activate the autophagic machinery in response to extreme mitochondrial
dysfunction and disruption in the metabolic machinery. A role for MCL-1 in the regulation of
mitochondrial bioenergetics was proposed via studies in Mcl-1-deficient fibroblasts that
displayed improper assembly of ATP-Synthase subunits [221]. Preliminary results have
supported this interaction as MCL-1 was immunoprecipitated with ATP Synthase and ATP
Synthase- assembly factor (ATPAF and ATP5B) in total ovarian lysates (data not shown). This
would indicate that the block in mitochondrial bioenergetics evident in Mcl-1cKO oocytes could
be due to the inability to assemble functional ATP-Synthase. Oocyte quality has long been
considered an essential factor in determining reproductive competency and has been suggested to
rely on mitochondrial content and effective mitochondrial bioenergetic output [112-115].
Maternal age has been correlated with increases in cellular oxidative damage, increased cellular
fragmentation, decreased metabolic output and elevated mitochondrial dysfunction [4-7].
Confirmatory studies are required to determine whether increased maternal age and impaired
oocyte quality are associated with Mcl-1 depletion.
We have shown that wildtype atretic follicles in vivo are characterized by a complete absence of
MCL-1 staining in the oocyte, indicating that down-regulation of MCL-1 could precede initiation
of oocyte atresia. Interestingly, rates of atresia in post-pubertal Mcl-1cKO ovaries are unaffected;
instead the lack of Mcl-1 seems to escalate the rapidity of death in those follicles already
predestined to die. Classical follicular atresia, which has been associated with concurrent
expression of markers of autophagy and apoptosis [142], is preceded by the death and recession
of surrounding granulosa cells [146, 147], in addition to the resumption of meiosis [260].
Inhibition of meiotic resumption, even during DNA damage-inducing death stimuli, prevents the
activation of apoptotic cellular fragmentation [261]. As oocyte-cumulus cell contact is required
90
for the maintenance of meiotic arrest [77-79], it is conceivable that with the maintenance of
granulosa/cumulus cell contact and support, the Mcl-1-deficient oocyte can activate autophagy,
but may be prevented from undergoing classical atresia.
Furthermore, the death or withdrawal of surrounding granulosa cells during follicle atresia [146,
147] has also been linked to starvation of the oocyte, as oocyte-cumulus cell contact has been
demonstrated to be essential for oocyte metabolic support [65, 81, 82, 94, 95]. Granulosa cells
have been shown to provide various metabolites, and possibly ATP, to the oocyte through gap
junctions maintained at regions of oocyte-granulosa contact. It is thus reasonable to assume that,
as granulosa cells regulate metabolite supply to the oocyte, any metabolic deficiency in Mcl-
1cKO GV oocytes may be overcome by a direct energy supply from surrounding granulosa. This
may account for the lack of reduction of total ATP output in freshly isolated oocytes from Mcl-1-
cKO ovaries, as compared to wildtype controls, despite significant reduction in downstream
TCA cycle substrates and respiring mitochondria number. Additionally, as follicle atresia is
granulosa cell-instigated, and the maintenance of granulosa cell contact can continue metabolic
support to the oocyte, this may also explain why Mcl-1cKO ovaries do not display an increase in
follicle atresia rates. Hence, upon commencement of granulosa cell death or withdrawal,
compromised Mcl-1cKO oocytes undergo a more rapid atretic demise, as exhibited by increased
counts of late-stage atretic follicles in post-pubertal ovaries.
These hypotheses are further supported by the finding that those Mcl-1cKO oocytes that survive
till ovulation (MII), display the same autophagic phenotype as Mcl-1cKO GV oocytes, in
addition to activation of the apoptotic pathway and an increased propensity to fragment once
removed from granulosa cell contact. Bax, which has been shown to be dispensable in follicle
atresia in young post-pubertal mice [161], appears uninvolved in the Mcl-1cKO GV phenotype,
91
as the reduction in actively respiring mitochondria is retained in Mcl-1c/BaxDKO oocytes.
However, additional studies must be conducted to verify whether Bax is truly not involved in
follicle atresia, or whether additional redundant pro-apoptotic Bcl-2 effectors substitute its
function. Additionally, as we do see an increase in BAX activation in Mcl-1cKO GV oocytes,
Bax may also play a novel non-apoptotic role in regulation of growing oocyte fate. Remarkably,
upon ovulation, Bax-deletion rescues the predilection of the compromised Mcl-1cKO denuded
MII oocytes to fragment in culture. As MII oocyte fragmentation has been classified as
containing hallmarks of apoptotic cell death [148], it is likely that Mcl-1cKO MII oocytes
undergo cellular fragmentation due to BAX activation. These results confirm recent findings
where Mcl-1-ablation was associated with the activation of autophagy in cortical neurons and
cardiac myocytes [203, 230]. Germain et al. claim that Mcl-1-deficiency alone is not sufficient
to activate cell death in cortical neurons [203]. Instead, in the absence of Mcl-1, it is the
activation of Beclin-1 that results in autophagy, followed by instigation of the apoptotic cascade
upon induction of pro-apoptotic Bcl-2 members. Additionally, the elevated mitochondrial
dysfunction associated with Mcl-1-ablation in cardiac myocytes, and proposed to be the
causative factor leading to the activation of autophagy, was unaltered with the concomitant
deletion of Bax and Bak [230]; further supporting the hypothesis that Mcl-1 plays an additional
role in mitochondrial function, possibly through the MCL-1Matrix
isoform. Thus, although Bax-
deficiency may rescue Mcl-1-deficient MII oocyte fragmentation in culture, the unrescuable
Mcl-1-deficient phenotype (mitochondrial dysfunction) most likely results in the compromised
developmental capacity of the resulting zygote. This postulation remains to be tested.
Fertility and reproductive proficiency has been well established to rely on the maintenance of the
ovarian reserve in addition to preservation of oocyte quality [3-7]. Despite the putative existence
of germline stem cells in the adult ovary [8], the conservation of fertility and the overcoming of
92
age-related factors governing oocyte competence remains extremely important in today’s
society, where a delay in child-bearing has become more of the norm [274]. Oocyte quality with
respect to maternal age has been associated with a variety of risks, including increases in
aneuploidies, cellular oxidative damage and cellular fragmentation, and decreased mitochondrial
function and metabolic output [4-7]. Interestingly, in this study we show that ablation of Mcl-1
results in the persistence of the majority of these factors associated with compromised oocyte-
quality. Essentially, we establish that Mcl-1 plays the defining role in mediation of oocyte
survival via protection of the postnatal primordial follicle pool, in addition to the growing oocyte
pool both prior to and upon ovulation. Mcl-1 regulates this survival via inhibition of factors
regulating PCD, and likely the management of mitochondrial functionality and output. This puts
Mcl-1 at the nexus of all efforts for maintenance of reproductive competency via preservation of
oocyte quality and survival.
93
3 CYTOKINE AND METABOLIC REGULATION OF MCL-1
FUNCTION IN MURINE OOCYTES
3.1 INTRODUCTION
Oocyte-granulosa intercommunication is vital for oocyte survival, proper follicular development
and dynamics, granulosa cell expansion, maintenance of oocyte meiotic arrest, in addition to the
provision of metabolite support for the oocyte (Elvin, Clark et al. 1999; Eppig 2001; Paulini and
Melo 2011). One of these important means of intercommunication includes the granulosa cell
secreted cytokine KL, which has been well established to bind the oocyte receptor tyrosine
kinase (C-KIT) and mediate aspects of survival and follicle dynamics [34, 36, 37, 40]. In
addition to ligand-receptor communication, Anderson et al. have identified the presence of gap
junctions of varying lengths between the granulosa and the oocyte [66]. The importance of these
gap junctions in maintaining oocyte meiotic arrest, follicle survival and proper follicular
development has been well demonstrated [70, 77-79]. Additionally, these gap junctions are
essential for the transfer of various metabolites to the oocyte, including amino acids, pyruvate,
cholesterol, and even ATP [65, 81, 82].
The oocyte itself seems incapable of full utilization of most means of energy breakdown. Key
experiments revealed that although cumulus-oocyte complexes (COC) were able to utilize
glucose, pyruvate, phosphoenolpyruvate or lactate, denuded oocytes were only able to
breakdown pyruvate or oxaloacetate [82, 83]. In fact, oocytes display a very few number of
glucose transporters and are able to survive and thrive in the absence of key glycolytic enzymes
94
[84-86]. Hence, oocytes are extremely reliant on granulosa cell support to provide key
metabolites required for oocyte mitochondrial bioenergetics. The TCA cycle and oxidative
phosphorylation, essential portions of aerobic respiration, both take place within the
mitochondria.
In addition to roles in cell metabolism and energy production, mitochondria are also the site of
the intrinsic pathway of apoptosis. The intrinsic pathway of apoptosis is mediated effectively by
the Bcl-2 family of pro- and anti-apoptotic factors that control cell fate [126-128]. One of these
Bcl-2 family members, the pro-survival factor Mcl-1 (Kozopas, Yang et al. 1993), was found to
be expressed in fetal human oocytes of the primordial follicle pool, oocytes of preantral follicles
and granulosa and theca cells of growing follicles [192, 193]. Our previous work has revealed
that murine oocyte-specific Mcl-1 ablation resulted in a sharp reduction of follicles of all stages
in neonates and triggered a POF-state by 4 months. Mcl-1-deficient oocytes recovered at 3 weeks
displayed increased markers of apoptosis, autophagy and mitochondrial dysfunction, albeit in a
stage-specific manner. Concomitant deletion of the pro-apoptotic Bcl-2 member Bax was able to
rescue follicle numbers in oocyte-specific Mcl-1-deficient mice, but dual Mcl-1 and Bax-
deficient oocytes continued to display increased markers of mitochondrial dysfunction and
autophagy. Ablation of Bax has been linked to increased primordial germ cell survival, an
increased primordial follicle pool, and extended fertility [156, 160], however Bax was shown to
be dispensable for follicular atresia [161].
In addition to mediation of the intrinsic pathway of apoptosis, select Bcl-2 members have been
implicated in the maintenance of cell metabolism. Bcl-xL has been attributed to regulation of
ATP Synthase stability, with increases in proton leakage apparent in Bcl-xL-ablated neurons
[220]. Furthermore, a mitochondrial matrix-specific isoform of MCL-1 (denoted as the MCL-
95
1Matrix
), the result of a mitochondrial processing peptidase (MPP)-mediated N-terminal cleavage,
may also have an ostensible role in ATP Synthase assembly and thus ATP production [221].
In this study, we explored the regulation of oocyte MCL-1 via two granulosa cell-mediated
events, downstream of growth factor induction and nutrient-starvation. We also describe the role
of MCL-1 in maintenance of mitochondrial bioenergetics in the growing follicle oocyte pool.
The KL cytokine-activated Class 1A Phosphotidylinositol 3 Kinase (PI3K) pathway appears to i
full-length MCL-1 protein expression via phosphorylation and inhibition of GSK-3 in pooled
primordial and primary oocytes. The granulosa cell-secreted ligand KL has been demonstrated to
activate the PI3 Kinase pathway in oocytes [34, 38, 40], a pathway that has been extensively
documented to regulate primordial follicle activation and survival [18, 22, 23, 29, 30, 33]; yet the
downstream molecules mediating follicle survival remain unidentified. The MCL-1Matrix
isoform
appears unchanged with modulation of the PI3 Kinase pathway; but does seem to be
metabolically regulated, as pyruvate-starvation leads to its depletion. These data suggest that
oocyte MCL-1 is growth factor-regulated and nutrient-sensitive; and a reduction in MCL-1,
through putative activity of the MCL-1Matrix
isoform, is linked to disruption of metabolic output,
thus leading to oocyte atresia.
In summary, we implicate Mcl-1 as an important crossroads for mediation of the growing oocyte
fate. Immunostaining of post-pubertal histological sections reveals the sharp reduction in oocyte
MCL-1 levels in growing follicles undergoing the first stages of atresia. Follicle atresia has long
been held as a granulosa cell-mediated event characterized by granulosa cell withdrawal [146,
147], and here we demonstrate two means of regulation of MCL-1 by granulosa cell controlled
events. Through the joint or mutually exclusive efforts of granulosa cell-secreted KL and the gap
junction-mediated supply of metabolites, granulosa cell contact can maintain MCL-1 levels and
96
hence oocyte survival. Conversely, down-regulation of MCL-1 in oocytes can effectively hasten
the granulosa cell-enforced atresia, permitting activation of autophagy and apoptosis, in addition
to a new role of inefficient mitochondrial machinery and disrupted metabolic output.
3.2 MATERIALS AND METHODS
3.2.1 Animals
For creation of a mouse line with an oocyte-specific deletion of Mcl-1, C57Bl6 Mcl-1tm3Sjk
(Mcl-
1f) [226] mice containing the floxed allele (which were a kind gift from the breeding colony of
Dr. Joseph T. Opferman) were intercrossed to mice carrying the Tg(ZP3Cre)3Mrt (Zp3-Cre)
transgene [263] (backcrossed from ICR/129 for 5 generations on to the C57BL/6 background)
generating the oocyte-specific Mcl-1t2Sjk
(Mcl-1null
) allele. Reporter lines utilized to assess timing
of Cre excision have already been discussed in the previous chapter. For early and complete
excision, Mcl-1f/null
: Zp3-Cre (hereafter referred to Mcl-1cKO) females were utilized in our
studies rather than Mcl-1f/f
: Zp3-Cre mice. Additionally, Mcl-1f/null
, Mcl-1+/+
, and Mcl-1f/+
: Zp3-
Cre females were collected as controls. Genotyping primers used for identification of Mcl-1+,
Mcl-1null
, Mcl-1f allele or Zp3-Cre transgene have also been documented in the previous chapter.
ICR mice were acquired from Toronto Centre for Phenogenomics (TCP) in-house breeding
colonies. All mice were housed with free access to food and water and were maintained on a
12h:12h light-dark cycle. All mouse experiments were performed in accordance with the
Canadian Council on Animal Care (CCAC) guidelines for Use of Animals in Research and
Laboratory Animal Care, under protocols approved by animal care committees at Mount Sinai
Hospital (MSH) or the TCP.
97
3.2.2 Collection of GV Oocytes
Mice from PN21-28 were primed with by stimulation with 10U PMSG (Pregnant Mare Serum
Gonadotropin; NHPP, USA or ProSpec, Israel (HOR-272)). PMSG-primed ovaries from mice
40-48hrs after stimulation were collected in mHTF and pierced with a small gauge needle
releasing antral COCs. Denuded oocytes were collected by manually stripping cumulus cells
from these COC’s using narrow bore glass pipettes. Diploid oocytes from the growing follicle
pool are arrested in diplotene stage of prophase I and characterized by a large nucleus termed the
germinal vesicle (GV).
3.2.3 Collection of Growing Oocyte Pool – PI3 Kinase Pathway
Representative oocyte samples of the growing oocyte pool were collected using a modified
protocol from Liu et al. [38]. Briefly, female wildtype ICR mice at PN8-10 were sacrificed and
ovaries were collected in Dulbeccos Modified Eagles Medium (DMEM)/F12 (Gibco-Life
Technologies) media supplemented with 4mg/ml Bovine Serum Albumin (BSA). These ovaries
were minced and incubated in the above media, with the addition of 500µg/ml collagenase for
45-60 min with constant pipetting and vortexing. 40mM EDTA was added and the mixture was
incubated at 37ºC for 15-20 min with constant pipetting and vortexing. This cell mixture was
then plated on a 60mm tissue culture dish with fresh medium and incubated for 6hrs allowing the
granulosa cells to attach to the surface of the dish. Floating oocytes were then collected,
centrifuged (1000rpm) and transferred to a 24-well plate. Oocytes were treated for designated
times with KL (SCF) 150ng/ml (Sigma), PI3 Kinase inhibitor LY294002 (Cayman Chemicals) at
98
25µM, GSK-3 inhibitor IX (Santa Cruz) at 200nM or vehicle (DMSO). Cells were collected for
western blot analyses.
3.2.4 Collection of Growing Oocyte Pool – Pyruvate Treatment
Oocytes from the growing oocyte pool were collected from wildtype ICR mice at PN14-18
utilizing a protocol similar to one described above. Ovaries were collected in glucose and
pyruvate free DMEM (Gibco-Life Technologies) supplemented with 4mg/ml BSA and 1mM
sodium pyruvate (Gibco-Life Technologies). Floating oocytes were transferred to 24 well plates
with treatments at designated times with or without 1mM pyruvate. Cells were collected for
western blotting.
3.2.5 Treatment with Inhibitors of Pyruvate Uptake and Fatty Acid Breakdown
GV oocytes from Mcl-1cKO and controls were collected in mHTF medium in the presence of
0.5mM 3-isobutyl-1-methylxanthine (IBMX) (Sigma), an inhibitor of meiotic progression. After
3 washes, oocytes were then incubated in 50µl droplets of HTF medium with IBMX, in the
presence of indicated inhibitors: 0.5mM α-hydroxycinnamic acid (Sigma), an inhibitor of
pyruvate transport to mitochondria; and 100µM etomoxir (Sigma), an inhibitor of carnitine
palmitoyl-transferase-1 (CPT-1), required for fatty acid transport to mitochondria; or vehicle
(DMSO). Oocytes were cultured for 96hrs, with survival rates measured at 8hrs, 16hrs, 24hrs,
36hrs, 48hrs, 72hrs and 96hrs. Viability of cultured oocytes was assessed using propidium
iodide/acridine orange stain.
99
3.2.6 Co-Immunoprecipitations
RIPA lysates were produced and collected from 30 PN4 ovaries, in addition to oocyte-enriched
assays from 40 PN18-21 animals as described in the growing oocyte pool collection protocol
above. PN4 ovaries represent a primordial-only pool of follicles, whereas the PN18-PN21
animals represented a growing pool of oocytes of all stages. 300µg of PN4 lysates or oocyte-
enriched lysates were pre-cleared in the presence of Protein A/Protein G sepharose bead mix,
shaking for 1hr at 4ºC. Lysates were spun down (1000g), beads removed and then incubated
overnight, shaking with 4µl (1mg/ml) of Mcl-1 Rockland antibody overnight at 4ºC. 10µl of
Protein A sepharose beads were added and incubated, shaking at 4ºC for 4hrs. Lysate-bead mix
was spun down (1000g), and heated at 95ºC for 3 min and flow through was collected. Beads
were washed 3 times in RIPA buffer, spinning down (1000g) between each. Lysates, washes and
flow through were run on a 12% SDS gel, and then transferred to a PVDF membrane. Blots were
incubated as described earlier, with ATP5β (Santa Cruz, sc-55597) and ATPAF (Santa Cruz, sc-
161370).
3.2.7 Western Blots, Antibodies, Reagents
The following antibodies were used for western blotting: Mcl-1 (Rockland Immunochemicals,
600-401-394S), Actin (Santa Cruz, sc-1616), p-Akt S473 (Cell Signaling, 9271S), Akt (Cell-
Signaling, 9272), Bax (Santa Cruz, sc-526), Bim (Millipore, MAB17001), Bim (Cell Signaling
C34C5), and GSK-3 Antibody Sampler Kit (Cell Signaling, 9369).
100
3.2.8 Measurement of ATP and Lipid Droplets
For quantification of ATP, single oocytes were collected for ATP analyses using CellGloTiter
Luminescent Cell Viability Assay (Promega). GV oocytes were collected in mHTF medium and
then incubated in HTF medium containing 50µM BODIPY 493/503 (Invitrogen) for 30minutes.
Oocytes were then washed 3 times in mHTF medium and imaged with LEICA DMI60003
Spinning disc confocal microscope. Lasers for the following wavelengths were used: DAPI-blue
(405 excitation, 450 emission), FITC-green (491 excitation, 525 emission). Representative
images included are high resolution de-convolved images.
3.2.9 Statistics
Statistical measures on two samples were performed using the unpaired T-test (pyruvate
treatment (wildtype GV oocytes)) or paired T-test (d8-14 culture (pyruvate/KL)); and additional
data was analyzed using two-way ANOVA (culture with inhibitors of pyruvate uptake (wildtype
and Mcl-1cKO GV oocytes), pyruvate and α-ketoglutarate treatment of wildtype and Mcl-1cKO
GV oocytes).
3.3 RESULTS
Mcl-1 is intricately involved in the regulation of oocyte survival and maintenance of the
primordial follicle pool, as has been established in the previous chapter. We observed MCL-1
expression in the cytoplasm of primordial follicle oocytes, and an increased accumulation of
MCL-1 from the primordial to primary transition and maintained with further follicle growth,
indicating that primordial follicle activation is associated with the upregulation of pro-survival
101
factors. In order to test this hypothesis, we decided to investigate whether MCL-1 could be
regulated by growth factors known to stimulate oocyte growth.
3.3.1 Impact of KL-Activated PI3 Kinase Pathway Stimulation on MCL-1
KL-activation of the PI3 Kinase pathway has previously been verified in a primary oocyte
culture model, resulting in phosphorylation and resultant inhibition of GSK-3 [38]. GSK-3 has
been acknowledged for roles in cell proliferation, cell migration, apoptosis and cell metabolism
in various cell types [39]. Importantly, its role in the phosphorylation and inhibition of MCL-1;
resulting in its reduced stability, increased degradation and impaired binding to pro-apoptotic
Bcl-2 members has also been documented, however appears to be cell type specific [222, 238,
275]. To determine whether KL-activation, or members of the PI3 Kinase pathway regulate
MCL-1 protein levels, through phosphorylation and inhibition of GSK-3 activity, oocytes
isolated from PN8-10 ovaries were cultured in the presence or absence of various
activators/inhibitors of the KL-directed PI3 Kinase pathway.
Treatment with the KL resulted in a significant elevation of the full length isoform of MCL-1
2hrs after treatment initiation, with no apparent change in mitochondrial MCL-1Matrix
isoform
levels (Fig. 3.1). This elevation was still evident 12hrs after treatment initiation (data not shown).
KL activation of the PI3 Kinase pathway was verified with increased phosphorylation of Protein
Kinase B (PKB)/AKT, and GSK-3. Concomitant treatment of KL-activated cultures with the PI3
Kinase inhibitor LY294002 significantly reduced full length MCL-1 twofold, with no change in
MCL-1Matrix
isoform (Fig. 3.1). LY294002 activity was validated by reduced phosphorylation of
AKT and GSK-3. Finally, treatment with the GSK-3-inhibitor IX significantly increased full
length MCL-1 levels twofold, with a small but significant increase in MCL-1Matrix
as well (Fig.
102
3.1). This rise in MCL-1Matrix
may be due to additional mechanisms of GSK-3 regulation of
MCL-1 expression or stability. In addition to its roles in cell proliferation, cell migration and
apoptosis, GSK-3 plays an essential part in the regulation of cell metabolism [39, 276-279].
3.3.2 Prevention of Oocyte Death with Pyruvate Supplementation
Recent findings by Perciavalle et al. have identified the MCL-1Matrix
isoform as a mitochondrial
matrix-specific cleavage product of the MPP [221]. The MCL-1Matrix
isoform is not involved in
regulation of the apoptotic machinery, implying that regulation of cell death is mediated by the
cytoplasmic/OMM -bound full length isoform. Mcl-1-deficient mouse embryonic fibroblasts also
displayed impaired ATP-Synthase assembly, which implies a putative metabolic role for MCL-1.
MCL-1 has also been shown to be sensitive to glucose-starvation in various in vitro models [203,
255], and we have verified disruption of TCA cycle substrates in Mcl-1cKO oocytes; thus we
hoped to determine the role of MCL-1, and especially the MCL-1Matrix
isoform, in the regulation
of oocyte metabolism.
Early studies on oocyte metabolism have established that the oocyte itself is unable to utilize all
avenues of energy production. COCs have proven capable of a wide range of metabolite
breakdown, but denuded oocytes only demonstrated breakdown of pyruvate or oxaloacetate [82,
83]; thus displaying a significant preference for aerobic respiration and hence oxidative
phosphorylation for energy production. Pyruvate utilization has been shown to increase with
growth, as does oxygen used for oxidative phosphorylation. Maximal pyruvate usage, when
controlling for volume, occurs in oocytes from primary follicles [92]. These experiments, in
addition to those performed by Biggers et al. and Eppig et al. designates aerobic respiration as
the prime means of energy production in the oocytes [82, 83]. As oocytes have been
103
Figure 3.1. Activation/Inhibition of PI3 Kinase Pathway and Impact on MCL-1 Expression.
The impact of inhibition or activation of the Kit ligand-activated PI3 Kinase pathway, associated with
oocyte survival, on MCL-1 expression. The growing follicle pool was isolated from day8-12 wildtype
ovaries, and cultured for 2hrs with KL (150ng/ml), LY294002 (25uM) or GSK-3 inhibitor IX (200nM).
(i) Image displaying pathway to be tested, with inhibitors and site of impact. (ii) Western Blots (WB)
showing impact of various culture treatments on MCL-1 full length, in addition to MCL-1Matrix
isoform.
ACTIN used as internal control. Each blot shows representative image of paired comparisons of each
treatment plus vehicle control on both MCL-1 isoform levels; specifically incubation with KL and vehicle
(n=5), KL with LY294002 (inhibitor of PI3Kinase activity) and KL with vehicle (n=4), and GSK-3
inhibitor and vehicle (n=4). Below, WBs of paired comparisons confirming the various treatments via
impact on phosphorylated AKT and phosphorylated GSK-3, both downstream members of the PI3 Kinase
pathway. (iii) Impact of the oocyte culture treatments using KL (purple), LY294002 (yellow) or GSK-3
inhibitor (green) on levels of MCL-1 full length isoform, band intensity quantitated with Quantity One
Software and normalized to ACTIN control. (iv) Impact of the same oocyte culture treatments on the
MCL-1Matrix
isoform, quantitated with Quantity One Software and normalized to ACTIN. Graph values
(iii and iv) represent average fold change of quantitated full length MCL-1 or MCL-1Matrix
± SEM with
treatment, normalized to vehicle-treated at same time-point. (*= p<0.05, ***= p<0.001).
104
demonstrated to take up pyruvate directly from their surroundings [82, 91, 280], we first verified
the importance of pyruvate in early oocyte survival. The growing oocyte pool (composed of
secondary and preantral follicles) was isolated from pre-pubescent (PN14-18) ovaries denuded of
granulosa cells and subsequently incubated for 12hrs in glucose-free DMEM medium with or
without 1mM pyruvate. Pyruvate starvation significantly increased the percentage of cell death
from 58% in the presence of pyruvate, to 90% once pyruvate was removed (Fig. 3.2A). In culture
conditions, denuded GV oocytess underwent cell death via oocyte lysis or shrinkage, rarely ever
by cellular fragmentation; which is reminiscent of autophagic or necrotic cell death and not
classical apoptosis.
To further validate the previous findings of the importance of pyruvate in post-pubertal denuded
antral follicles [82, 83], we also isolated PMSG-stimulated antral follicle oocytes and incubated
them for 12hrs in the same medium utilized above, in the presence and absence of pyruvate (Fig.
3.2A). Supplementation with pyruvate provided a distinct survival advantage as the percentage
of dead GV stage oocytes were significantly increased from 5% with pyruvate to 65% upon
starvation.
3.3.3 Importance of Mcl-1 in Oocyte Metabolism.
We have previously demonstrated that follicles undergoing atresia in wildtype ovaries strongly
down-regulate MCL-1 expression. This implies that regulation of MCL-1 is strongly correlated
with the advent of follicle death. In order to assess whether down-regulation of MCL-1 in
oocytes precedes initiation of follicle atresia, and to establish that the MCL-1 response mediates
the increase in oocyte survival, we performed WBs on oocyte-enriched pools of PN14-18
105
ovaries, cultured for 6 hours with or without metabolite support. Oocytes cultured without
pyruvate, exhibited a significant decrease in levels of the MCL-1Matrix
isoform, when compared
to pyruvate-supplemented controls, with a smaller but significant decrease in full length MCL-1
levels as well (Fig. 3.2B). This confirms previous findings that demonstrate sensitivity of MCL-1
to nutrient deprivation in vitro [203, 255], and also establishes oocyte sensitivity to nutrient
deprivation. In various in vitro cell models, (cortical neurons, B- and T- cells) nutrient
deprivation has been associated with a reduction in MCL-1 levels, and subsequent activation of
pro-apoptotic Bcl-2 members and increased cell death [203, 255, 281]. To determine whether
MCL-1 plays a similar role in oocyte survival in the absence of nutrients, we cultured denuded
Mcl-1cKO GV oocytes and wildtype controls in the presence or absence of pyruvate or α-
ketoglutarate. α-ketoglutarate is a crucial substrate of the TCA cycle, downstream of pyruvate
metabolism. As expected, in control GV oocytes cultured for 12hrs, pyruvate or α-ketoglutarate
starvation significantly increased death rates to 65% from around 8% and 12% in the presence of
pyruvate or α-ketoglutarate, respectively (Fig. 3.3A). Incubation of Mcl-1cKO GV oocytes with
pyruvate or α-ketoglutarate supplementation resulted in death rates comparable to controls,
however under nutrient-deprived conditions, Mcl-1cKO exhibited exacerbated death rates of
95% compared to 65% in controls (Fig. 3.3A). Further confirmation of the increased
susceptibility to cell death of the Mcl-1cKO GV oocytes is that the survival associated with α-
ketoglutarate supplementation was not maintained, as by 24hrs, 100% of Mcl-1cKO oocytes
supplemented with α-ketoglutarate died, compared to 50% of wildtype controls (data not shown).
The delayed/reduced death of Mcl-1cKO oocytes in the presence of abundant metabolites,
suggests that either nutrient supplementation activates an additional redundant survival pathway
independent of MCL-1, inhibits activation of apoptotic pathways, or
106
Figure 3.2. Impact of Pyruvate Treatment on Oocyte Survival and MCL-1 Expression.
(A) The impact of pyruvate starvation on oocyte survival. Growing follicle pool oocytes from wildtype
pre-pubertal (PN14-PN18) animals were isolated and incubated in media with 1mM pyruvate (light blue)
or without (dark blue) for 12hrs. (i) Image displays growing follicle oocytes with both treatments, stained
with propidium iodide/acridine orange. Propidium iodide (orange) is a nuclear stain that detects apoptotic
cells, whereas acridine orange (green) detects all nucleated cells, both live and dead (ii) Oocytes from
n=3 experiments of 12hrs culture treatments with 1mM pyruvate (n=169) or starved (n=142) were
quantitated and graphed and values represent percentage of dead oocytes/total oocytes per experiment ±
SEM. (iii) GV oocytes were isolated from wildtype (PN21) females, stripped of granulosa cells and
subjected to 12hrs culture with 1mM pyruvate (n=3) and starved (n=3). Total GV oocytes used for
experiments were n=55 with 1mM pyruvate, and n=38 starved. Values represent percentage of dead GV
oocytes/total cultured GV oocyte pool per experiment ± SEM. (B) Impact of pyruvate starvation on
MCL-1 expression. The growing follicle pool was isolated and incubated with or without 1mM pyruvate
for 6hrs and Western blots (WB) performed to determine impact on full length MCL-1 and MCL-1Matrix
.
(i) The WB displays a representative image from n=5 experiments of paired comparisons performed
utilizing this culture treatment. Bands displaying impact on MCL-1 full length and MCL-1Matrix
are
presented with ACTIN used as an internal control. (ii) WBs were quantitated using Quantity One
Software and band intensity of MCL-1 full length (n=4) was normalized to ACTIN controls. (iii) Band
intensity of WBs for MCL-1Matrix
(n=5) were quantitated and normalized to ACTIN control. Values from
(ii and iii) represent fold change of quantitated full length MCL-1 or MCL-1Matrix
± SEM per treatment,
normalized to pyruvate treated at the same time-point. Analysis was performed using paired T-test. (*=
p<0.05)
107
108
Figure 3.3. Impact of Starvation on Oocyte Survival in Mcl-1cKO.
(A) GV oocytes were isolated from young PN21 mice, stripped of granulosa cells and cultured in various
treatments for 12hrs. Oocyte death was calculated from Mcl-1f/null
: Zp3-Cre (Mcl-1cKO) (red) (n=3, 3, 4)
and Mcl-1+/+
(blue) (n=4, 3, 3) in 1mM pyruvate, 5mM α-ketoglutarate or starved culture conditions
respectively. The total number of oocytes utilized from Mcl-1cKO females were n=34 in starved
conditions, n=25 in media supplemented with pyruvate and n=26 in media with α-ketoglutarate; whereas
total oocytes from Mcl-1+/+
females were n=36 in starved conditions, n=55 with pyruvate and n=44 in α-
ketoglutarate. Graph values represent percentage of dead GV oocytes/total GV pool ± SEM per
experiment. Statistical analysis reveals significant difference in comparison of pyruvate vs starved
(p<0.001) or α-ketoglutarate vs starved (p<0.001) in Mcl-1cKO and in in comparison of pyruvate vs
starved (p<0.001) or α-ketoglutarate vs starved (p<0.001) in Mcl-1+/+
. Also in comparison of Mcl-1cKO
vs Mcl-1+/+
in starved conditions (p<0.05) (B) GV oocytes, isolated from PN21 mice and stripped of
granulosa cells were incubated in HTF medium with 0.5mM IBMX to prevent meiotic progression. GV
oocytes were treated with 0.5mM α-hydroxycinnamic acid (α-HCA), an inhibitor of mitochondrial
pyruvate uptake, or vehicle; and oocyte death was documented at indicated timepoints. Culture treatments
were performed on a total of n=3 experiments for Mcl-1cKO and Mcl-1f/-
GV oocytes and n=4 for Mcl-
1+/+
. The total number of oocytes utilized in these experiments were n=38 with vehicle and n=39 with α-
HCA for Mcl-1cKO; n=27 with vehicle and n=30 with α-HCA for Mcl-1+/+
; and n=28 with vehicle and
n=27 with α-HCA for Mcl-1f/-
. Values represent percentage of dead GV oocyte/total GV pool ± SEM per
experiment. Statistical analysis reveals significant difference in comparison of Mcl-1cKO to Mcl-1+/+
(p<0.01), Mcl-1cKO to Mcl-1f/-
(p<0.05) and Mcl-1cKO 16hrs to 24hrs (p<0.001). Datasets marked with
same letter (a, b or c) indicate no significant difference between datasets, different letters indicate
significant difference.
109
MCL-1 is involved in maintenance of metabolic output (mitochondrial bioenergetics), and
treatment with abundant metabolites delays Mcl-1-deficient death. The seemingly transient
rescue of Mcl-1cKO oocytes with α-ketoglutarate supports the latter.
3.3.3 Regulation of Energy Output in Mcl-1-deficient oocytes
To further confirm the compromised metabolic capacity of Mcl-1cKO GV oocytes, we attempted
to inhibit metabolite uptake by the mitochondria, essentially further starving the oocyte. Denuded
Mcl-1cKO GV oocytes and controls were isolated and cultured in HTF medium, containing an
ample supply of metabolites (glucose, 1mM pyruvate, lactate). As oocytes have been
demonstrated to utilize additional means of energy production after GV breakdown, through
lipid breakdown and β-oxidation of fatty acids [88, 89], we maintained meiotic arrest with
IBMX. Mcl-1cKO oocytes and controls were treated with α-hydroxycinnamic acid, an inhibitor
of mitochondrial pyruvate uptake [282]. Inhibition of pyruvate uptake resulted in complete
oocyte death after 48 hours in both control and Mcl-1cKO oocytes. However, Mcl-1cKO oocytes
showed an increased sensitivity to this inhibition, with 56% of oocytes dying by 24 hours of
culture compared to only 12-20% of control oocytes (Fig. 3.3B). This result displays an
increased susceptibility of Mcl-1cKO GV oocytes to cell death in the absence of pyruvate
metabolism. Taken together with the previous results it indicates a compromised metabolic
capacity of Mcl-1cKO GV oocytes, resulting in an elevated death potential when metabolites are
withdrawn. Furthermore, the short-term survival advantage provided by α-ketoglutarate
demonstrates an inefficiency in adequate utilization of the available metabolites by Mcl-1cKO
oocytes compared to controls, likely due to increased malfunction in the oxidative
phosphorylation machinery. The proposal for MCL-1 regulation of donwnstream metabolic
110
components is not a novel one, having been suggested by Perciavalle et al. who demonstrated
that Mcl-1KO MEFs display impaired ATP-Synthase assembly [221], implying a supplementary
metabolic role of MCL-1.
Additionally, BCL-x has been shown to play a role in mediation of efficient ATP-Synthase
function [220]. We attempted to verify the postulated interaction of MCL-1 with ATP Synthase,
in oocytes using co-immunoprecipitations (co-IPs). Preliminary MCL-1 co-IPs revealed that
MCL-1 did complex with ATP-5β, a subunit of ATP-Synthase, in addition to ATP-Synthase
Assembly Factor (ATPAF) (Fig. 3.4). Interestingly, MCL-1 co-IPs of total ovarian lysates from
neonatal animals, utilized as a representative of an enriched pool of primordial follicle oocytes,
did not complex with either ATP-5β or ATPAF (Fig. 3.4). This implies that the interaction of
MCL-1 and ATP-Synthase, and the alleged impact on ATP-Synthase assembly, does not occur
until after primordial follicle activation and subsequent growth. Further work using ATP-
5β/ATPAF pulldown (reverse pulldown) is required to ensure the specificity of this interaction.
As we have previously described, Mcl-1cKO MII ovulated oocytes assayed for metabolic
components of the TCA cycle and total ATP revealed no change in total ATP or citrate levels,
but a significant reduction in fumarate and malate (Fig. 2.6B). This evidence, in addition to data
from the previous chapter revealing the association of oocyte-specific Mcl-1-deletion with the
activation of autophagy, a starvation-induced process, and increased ROS, indicative of
overworked mitochondrial machinery, supported the role for MCL-1 in maintenance of
mitochondrial bioenergetics.
111
Figure 3.4. Co-Immunoprecipitation with MCL-1 Pulldown in Ovarian Lysates.
(i) Co-IP of total ovarian lysates of 3 week (PN21) females, using MCL-1 pulldown. Lysates were
immuno-precipitated with anti-Mcl-1 antibody, run on 12% SDS-PAGE gel and then transferred to PVDF
membrane. Blots were immuno-blotted with ATP Synthase subunit ATP5β, in addition to ATP Synthase
assembly factor (ATPAF); and with BH3-only activator BIM and pro-apoptotic Bcl-2 effector BAX. To
assess whether complex interaction was due to pulldown of outer mitochondrial membrane, co-IP was
immuno-blotted with translocase of outer mitochondrial membrane 20 (TOM20), a mitochondrial import
receptor. (ii) Co-IP of Day 4 (PN4) ovarian lysates using MCL-1 pulldown. PN4 ovaries are used as
representative of a large source of primordial follicles. Similar to the previous blot, these Co-IP blots were
immuno-blotted for interaction with ATP Synthase (using ATP5β and ATPAF) and pro-apoptotic Bcl-2
members (BAX and BIM).
112
3.3.4 Alternative Means of Energy Production
In addition to pyrvuate breakdown, mature oocytes and early zygotes have been documented as
utilizing lipid stores for energy supply [89]. Triglycerides make up the majority constituent of
the lipid store, and these triglycerides are broken down during lipolysis to supply fatty acids. β-
oxidation is the process via which the fatty acids are transported to the mitochondria where they
are degraded to provide acetyl coA stores for the TCA cycle. Although lipid storage and
breakdown varies between species, in mouse oocytes, the majority of lipid synthesis occurs after
GV maturation, both in vitro and in vivo [88]. Therefore, to confirm this finding, and to
determine whether Mcl-1cKO oocytes were utilizing lipid breakdown as an additional means of
energy production, we stained denuded Mcl-1cKO oocytes and controls with BODIPY, a
fluorescent marker of lipid droplets [283], at various developmental stages. GV oocytes of all
genotypes did not display much, if any lipid droplets; however lipid storage increased
tremendously in all genotypes upon GV breakdown (Fig. 3.5A). Freshly isolated MII oocytes
from all genotypes also exhibited varying degrees of lipid stores (Fig. 3.5A); however there was
no apparent change among studied genotypes.
To assess prolonged lack of granulosa cell contact on energy utilization via lipid storage, and to
determine whether supplementation with pyruvate or α-ketoglutarate would modulate lipid
storage capabilities, we matured denuded Mcl-1cKO and control GV oocytes in vitro for 6hrs in
the absence and presence of pyruvate or α-ketoglutarate. A proportion (76%) of control GV
oocytes cultured in the absence of pyruvate did undergo GV breakdown and subsequently
displayed an accumulation of lipid droplets (Fig. 3.5B). A comparable proportion (81%) of those
cultured in the presence of pyruvate also underwent GV breakdown, with the production of lipid
droplets of various sizes in 69% of the matured number. The entire population (100%) of control
113
Figure 3.5. Lipid Droplet Formation in Mcl-1cKO and Controls.
(A) Lipid content in freshly isolated GV and MII oocytes. GV oocytes and MII oocytes isolated from
stimulated PN21 females were denuded and stained with 50mM BODIPY (green), a fluorescent marker of
lipid droplets and counterstained with nuclear DAPI (blue). Images displayed are representative samples
of variable lipid droplet stores in Mcl-1f/null
: Zp3-Cre (Mcl-1cKO) freshly isolated GVs (a), with
brightfield image (f), oocytes that underwent GV breakdown (i.e resumption of meiosis) (b, c), with
brightfield image (g, h), and freshly isolated MII oocyte (d, e), with brightfield image (i, j). Also
displayed are lipid content of Mcl-1+/+
GVs (k), oocytes that underwent GV breakdown (l, m), and MII
oocytes (n, o); and Mcl-1f/-
GVs (p), GV breakdown (q, r), and MIIs (s, t). (B) Lipid content from freshly
isolated, denuded GV oocytes cultured for 6hrs in the presence of 1mM pyruvate, 5mM α-ketoglutarate or
starved. Representative images of Mcl-1cKO GV oocytes stained with BODIPY (green) and
counterstained with DAPI, after culture in starved (vehicle) conditions (n=7) (a, h), with pyruvate
supplementation (n=7) (b, i), or α-ketoglutarate supplementation (n=11) (c, j). Also displayed, in
comparison to lipid droplet content of Mcl-1cKO GVs, are representative images from Mcl-1+/+
GV
oocytes similarly stained after 6hr culture in starved (vehicle) conditions (n=17) (d, k), with pyruvate
supplementation (n=13) (e, f, l, m), or α-ketoglutarate supplementation (n=9) (g, n).
114
GV oocytes cultured in the presence of α-ketoglutarate displayed a tremendously strong increase
in lipid storage, and none of them exhibited signs of GV breakdown (Fig. 3.5B).
Conversely, the entire population (100%) of denuded Mcl-1cKO GV oocytes cultured for 6 hours
in the absence of pyruvate did undergo GV breakdown, but failed to accumulate any lipid stores
(Fig. 3.5B). This irregular in vitro maturation, accompanied by a lack of lipid droplet formation,
also occurred in all of the Mcl-1cKO GV oocytes matured in the media supplemented with
pyruvate. Moreover, treatment with α-ketoglutarate that, in controls, had resulted in a dramatic
increase in lipid storage, also failed to create any lipid stores in Mcl-1cKO GV oocytes (Fig.
3.4B). Remarkably, akin to control GV oocytes cultured with α-ketoglutarate, all of these
oocytes cultured with α-ketoglutarate displayed an inability to undergo GV breakdown.
The lack of lipid accumulation in cultured denuded Mcl-1cKO GV oocytes most likely implies a
lack of excess stores in the form of fatty acids, due to an increase in their utilization. It also
supports the idea of an over-utilization of all forms of metabolites for energy production, as we
postulated earlier. To determine whether fatty acid β-oxidation is an essential element in energy
production for GV oocyte survival, we incubated denuded Mcl-1cKO GV oocytes and controls in
HTF medium with IBMX, in the presence of Etomoxir, a compound that inhibits fatty acid
transport to the mitochondria. In the presence of adequate pyruvate and additional metabolites in
HTF medium, Mcl-1cKO GV oocytes exhibited no increased sensitivity to inhibition of fatty
acid β-oxidation compared to untreated oocytes as late as 72hrs of treatment (data not shown).
This suggests that pyruvate breakdown may be the preferred means of energy production in GV
oocytes, with fatty acid β-oxidation playing a supportive role in times of need; or a CPT-1-
independent (Etomoxir-impervious) mechanism may exist, promoting lipid uptake by
mitochondria. Interestingly, treatment with Etomoxir in conjunction with α-hydroxy cinnamic
115
acid, had an exaggerated effect irrespective of genotype, with the majority (>95%) of oocytes
dying as early as 36hrs (data not shown). Further analysis, and additional time-points may be
necessary to uncover any increased sensitivity of Mcl-1cKO GV oocytes in this scenario.
3.4 DISCUSSION
Like many other Bcl-2 members, Mcl-1 has been established to have additional roles aside the
classical roles in the inhibition of cell death. MCL-1 has been demonstrated to regulate cell fate
decisions of either autophagy or apoptosis [203, 219, 226, 227, 237, 254], in addition to roles in
cell cycle regulation and mediation of DNA repair [272, 273]. Studies outlined in this chapter
have focused on cytokine regulation of MCL-1, specifically by the KL-activated PI3 Kinase
pathway, as well as the roles of MCL-1 in metabolism. The outcomes of these oocyte studies
support recently described phenotypes caused by Mcl-1-deficiency in MEF’s [221]. Work by
Coloff et al. has revealed the dual role of MCL-1 in regulation of both growth factor and
metabolism-mediated cell survival in murine cell lines [255]. They demonstrate that removal of
either glucose or IL3 growth factor results in the reduction of MCL-1 levels; but expression of a
constitutively active AKT was only able to restore MCL-1 levels in a glucose-dependent manner.
In this chapter we have concentrated on the non-traditional (non-apoptotic/autophagic) roles of
Mcl-1, focusing specifically on the exogenous cytokine and metabolic regulation of MCL-1 in
oocytes, and the additional downstream role of the MCL-1Matrix
isoform in maintenance of
mitochondrial function and metabolic output. Data presented here, can help us further understand
somatic/granulosa cell regulated mechanisms mediating oocyte survival via regulation of MCL-
1, effectively controlling the induction of follicle atresia. Follicle atresia has been well
116
documented to be granulosa cell-mediated event involving the retraction of granulosa cells,
through death or withdrawal, leading to the effective cytokine and metabolic isolation of the
oocyte [146, 147]. As follicle atresia has been classified as bearing markers of both apoptosis
and autophagy [142], and full length MCL-1 has been associated with mediation of both
pathways [126-128, 203, 204, 219], it is likely that MCL-1 depletion precedes the atresia. In the
previous chapter, we have already demonstrated that wildtype atretic follicles, in vivo, are
characterized by a complete absence of MCL-1 staining in the oocyte, further solidifying the
association of MCL-1 depletion with eventual follicular demise.
With regards to exogenous cytokine modulation of oocyte fate, we have verified previous
findings that activation of the PI3 Kinase pathway by the cytokine KL, results in phosphorylation
of GSK-3 in cultured ex-vivo growing oocyte pools [38]. Taking this one step further we have
also demonstrated that oocyte expression of MCL-1 protein is also regulated by the KL-triggered
PI3 Kinase pathway in our ex-vivo primary oocyte culture model. Interestingly, PI3 Kinase
pathway activation results in elevation of the full length isoform of MCL-1, with no apparent
change in the MCL-1Matrix
isoform; which indicates that KL signaling may facilitate oocyte fate
via PI3 Kinase-mediated stabilization of full length MCL-1 protein. As the MCL-1Matrix
isoform
has been established to be uninvolved in regulation of cellular apoptosis, strictly restricting its
activity to mitochondrial functional maintenance [221], we hypothesize that cytokine regulation
of MCL-1 represents an exogenous (granulosa cell-directed) means of mediation of oocyte
MCL-1 activity and its associated impacts on apoptosis, autophagy and even mitochondrial
functionality and output. GSK-3, with varied roles in proliferation, survival and metabolism [39],
appears to regulate both the full length and MCL-1Matrix
isoform, implying additional PI3 Kinase-
independent interactions with MCL-1. GSK-3 phosphorylation of MCL-1, which can be
inhibited by activation of the PI3 Kinase pathway, has been linked to decreased MCL-1 protein
117
stability [222, 223]. Although we have assessed activation or inhibition of the PI3 Kinase
pathway at various timepoints, thus supporting a post-translational means of control of MCL-1
protein stability; transcriptional or translational regulation of MCL-1 by additional growth
factors, including KL [231, 235] is still a possibility that needs to be examined.
In addition to cytokine activation of MCL-1, we have also evaluated the metabolic role of MCL-
1 and putative duty of the MCL-1Matrix
isoform. The advent of follicle atresia is characterized by
granulosa cell death and withdrawal [146, 147] essentially starving the oocyte; as granulosa cells
are a major source of metabolites (pyruvate, amino acids, lactate) and perhaps even ATP,
transferred to the oocyte through gap junctions [65, 81, 82]. Thus in vivo oocyte starvation
appears to impact oocyte cell fate via regulation of MCL-1. To further test this metabolic angle,
we employed an in vitro nutrient/starvation model utilizing vehicle (DMSO), pyruvate or α-
ketoglutarate treatments, as denuded oocytes have been well documented to be solely dependent
on pyruvate or oxaloacetate breakdown for aerobic respiration and oxidative phosphorylation
[82, 83]. We confirmed this dependence as removal of pyruvate considerably decreased denuded
oocyte survival, and was associated with a significant reduction in MCL-1 protein levels,
specifically those of the MCL-1Matrix
isoform. This supports previous in vitro models, which have
established a nutrient-sensitive (glucose-dependent) down-regulation of MCL-1, with co-incident
increases in cell death [255, 281]. Further studies are required to identify the metabolically
sensitive factors regulating MCL-1 levels.
Intriguingly, although Mcl-1cKO GV oocytes did display increased cell death compared to
controls in the absence of nutrients and Mcl-1cKO GV oocytes were significantly more
susceptible to cell death when cultured with inhibitors of mitochondrial pyruvate transport;
pyruvate or α-ketoglutarate supplementation was able to delay Mcl-1cKO oocyte death. This
118
implies that with oocyte-specific Mcl-1-ablation, the machinery (aerobic respiration) utilized for
pyruvate or α-ketoglutarate breakdown remains functional; but fully competent bioenergetic
output may be impaired. However, it remains a possibility that the oocyte activates additional
means for metabolite breakdown, independent of MCL-1. Conversely, as glucose-starvation in
various in vitro models have been linked to the induction of pro-apoptotic factors [255, 281,
284], Mcl-1cKO GV oocytes, removed from the metabolic influence of granulosa cells, may also
be further susceptible to autophagic, necrotic or apoptotic cell death in nutrient-deprived
conditions.
There exists increasing amounts of evidence hinting at the possibility that MCL-1, most likely
the matrix-restricted MCL-1Matrix
isoform, is involved in maintenance of the mitochondrial
machinery. Perciavalle et al. initially postulated a role for the MCL-1Matrix
isoform in effective
ATP Synthase assembly and hence mediation of mitochondrial bioenergetics in MEF’s [221].
This notion was further supported by the defective mitochondrial phenotype associated with Mcl-
1-deletion in cardiomyocytes [230] and our findings in oocytes; where MCL-1 co-IP’s in oocyte-
enriched lysates reveal an interaction with ATP Synthase subunit ATP5β, and ATP Synthase
Assembly Factor (ATPAF). Additional work is required to identify whether this interaction is
direct or indirect, and what particular role MCL-1Matrix
may be playing in effective bioenergetic
output. Although Mcl-1cKO MII oocytes displayed no change in citrate or total ATP levels, a
disruption of mitochondrial bioenergetics is supported by the reduction of respiring
mitochondrial number, and the halving of total levels of Fumarate and Malate, constituents of the
TCA cycle downstream of α-ketoglutarate, observed in Mcl-1cKO oocytes.
Mitochondrial dysfunction associated with a defect in ATP Synthase assembly would thus give
rise to the faulty mitochondrial machinery as mentioned earlier. Additional supplies of
119
metabolites would be utilized, but with lower energy output, hence resulting in a constitutive
over-utilization of the mitochondrial machinery to supply the cell with adequate energy. This
putative over-utilization due to Mcl-1-ablation is supported by a number of lines of evidence. In
the previous chapter, Mcl-1cKO GV oocytes have been shown to activate macroautophagy, a
process by which cellular organelles are degraded for nutrient supply, utilizing the lysosomal
machinery of the cell, in order to maintain a consistent supply of energy [196, 285]. Elevated
mitochondrial dysfunction due to Mcl-1 loss has previously been postulated to play a causative
role in the activation of autophagy in cortical neurons and cardiac myocytes [203, 230]. As
described in the previous chapter, Mcl-1cKO oocytes also display increases in mitochondrial
dysfunction, associated with an elevation in total mitochondria, yet significant reduction in
actively respiring mitochondria, in addition to elevated ROS levels. Elevated ROS levels have
also been postulated as a downstream effect indicative of the overuse of the mitochondrial
machinery [119, 120]. Finally, culture of denuded Mcl-1cKO GV oocytes severely impacts
energy storage, as despite the presence of adequate metabolites, Mcl-1cKO GV oocytes fail to
create excess lipid stores upon maturation. These data altogether strongly suggest that MCL-1,
likely the MCL-1Matrix
isoform, is essential for normal mitochondrial function via efficient ATP-
Synthase assembly; and in the absence of adequate MCL-1, malfunctioning ATP-Synthase
results in the over-utilization of available metabolites and eventual cellular demise.
One interesting phenomenon we observed during oocyte culture of GV oocytes was that treat-
ment with α-ketoglutarate somehow inhibited GV breakdown, yet still resulted in the production
of excessively large lipid stores. As mentioned earlier, gaining the ability to synthesize lipid
stores in mouse oocytes has been associated with GV breakdown, meiotic progression and
oocyte maturation [88, 89], which we verified in culture systems treated with pyruvate or
vehicle. One possible explanation for this intriguing phenotype is that treatment with α-
120
ketoglutarate excessively stimulates production of downstream TCA cycle components, as the
conversion of α-ketoglutarate to succinyl coA is an irreversible step. This presumably leads to
excessive buildup of upstream citrate levels, resulting in the activation of fatty acid synthesis and
lipogenesis which may explain the excessive buildup of lipid stores in control GV oocytes. One
of the essential steps of fatty acid synthesis includes the conversion of malonyl coA to palmitate,
and malonyl coA has previously been attributed to inhibition of meiotic induction in GV oocytes
[286]. The extraordinary finding that Mcl-1cKO GV oocytes cultured with α-ketoglutarate do not
create lipid stores and still do not undergo GV breakdown, may be a result of transitional
quantities of malonyl coA preventing meiotic induction, or an additional novel impact of α-
ketoglutarate treatment on GV oocytes.
Oocyte quality is an extremely important determinant of reproductive function, as a reduction in
oocyte quality, accompanying advanced maternal age has been associated with a number of
developmental defects. These include increases in aneuploidies, decreases in DNA repair,
increases in cellular oxidative damage and cellular fragmentation, as well as impaired
mitochondrial function with decreased ROS and ATP production [4-7, 119, 120, 262]. Many
studies have linked the reduction in mitochondrial ATP output with spindle defects and
chromosomal abnormalities, leading to impaired zygotic development [116-118]. Through
understanding of the in vivo mechanisms utilized to induce oocyte death and follicle atresia, i.e
the two-pronged (granulosa cell-derived components) cytokine and metabolic reduction of MCL-
1, followed by the mitochondrial metabolic starvation induced by MCL-1Matrix
reduction; we may
be able to utilize the same means to boost mitochondrial bioenergetics, increasing oocyte quality
and preventing compromised reproductive outcomes.
121
4 OVERALL DISCUSSION
According to recent estimates, the mean child-bearing age for first births has increased
significantly from 1970-2006 in developed nations world-wide, and amongst these statistics, the
fraction of first births to women over 35 years of age has increased almost eight-fold [274].
Many studies have determined that the majority of the follicles of the ovarian reserve are lost to
atresia, with an exponential increase in follicular demise after 37 years of age, eventually
resulting in total reproductive senescence by menopause at a median age of 51 years [287, 288].
Moreover, decreases in oocyte quality have been linked to increased maternal age, exhibiting
high numbers of chromosomal abnormalities, ineffective DNA repair, mitochondrial dysfunction
and increased susceptibility to cell death [4-7, 289, 290]. Additionally, genetics, iatrogenic or
environmental factors can lead to a premature exhaustion of the follicle pool, resulting in a
condition termed POF [3]. Thus, understanding molecular pathways that govern the long-term
maintenance of the ovarian reserve is essential for developing strategies to prevent excessive
follicle loss.
Current methods of preservation of the primordial follicle pool, due to increased primordial
follicle loss induced upon chemotherapy or radiation treatments, is via cryo-preservation of
ovarian material or if possible, ovulated oocytes [291]. However, the feasibility of this option is
limited as the ability to mature follicles in in vitro cultures has been achieved in mice [292, 293],
but not yet been perfected in humans [294]. Oocyte vitrification has also been utilized to
preserve ovulated MII oocytes, with the hope to thaw them when required for In Vitro
Fertilization (IVF) [295]. Finally, mitochondrial nutrients or caloric restriction (CR) have been
utilized to improve oocyte quality in aged female mice, restoring them to a reproductive
phenotype more reminiscent of younger females [296, 297], and thus show promise for human
122
treatments. Determinants of oocyte quality and oocyte survival thus prove to be highly essential
to contend with situations of premature follicular exhaustion, and preservation of oocyte quality
to assist current infertility treatment protocols. We believe that one of those determinants is the
anti-apoptotic Bcl-2 family member Mcl-1.
Work described in the first study of this thesis characterized the stage-specific expression and
function of MCL-1 in mouse oocytes of neonatal and post-pubertal ovaries. We observed
transitional nuclear expression in primordial follicle oocytes of neonatal mice (Fig. 2.1A) which
disappeared in PN12 ovaries (data not shown). Cytoplasmic expression of MCL-1, more in
keeping with its known anti-autophagic and anti-apoptotic mitochondrial localization, was first
detected in primordial follicle oocytes, and was retained in the cytoplasm of growing follicle
oocytes. An increased aggregation of the cytoplasmic signal was associated with follicle growth,
implying that MCL-1 plays an important role in growing follicle dynamics.
A variety of anti-apoptotic Bcl-2 members have been found expressed in oocytes and granulosa
of growing follicles, however they have been found to play either a redundant or inessential role
in regulation of postnatal oocyte survival. Transgenic over-expression of Bcl-2 in neonatal
oocytes led to a fleeting mild increase in primordial follicle survival [180], whereas granulosa
cell over-expression was linked with a more operative survival phenotype [183]. Another anti-
apoptotic Bcl-2 member, BCL-x (Bcl2l1), was also identified in post-pubertal oocytes and
granulosa cells [184], a hypomorphic allele of Bcl-x was associated with embryonic loss of
primordial germ cells [158], but postnatal conditional deletion revealed no apparent impact on
postnatal germ cell survival [187]. Granulosa cell excision of Bcl-x revealed an apparently
redundant function and no phenotype. Finally, Diva (Bcl2l10) was localized to granulosa cells
and GV oocytes [188, 189], but total ablation of Diva or Bcl-w (Bcl2l2), revealed no ovarian
123
phenotype [190, 191]. Therefore, the phenotype associated with conditional oocyte-specific
deletion of Mcl-1, is the first described phenotype associated with a non-redundant function in
preservation of the post-natal ovarian reserve.
4.1 Regulation of Primordial Follicle Fate
Use of the Zp3-Cre transgene permitted Mcl-1-excision in oocytes as early as 17.5dpc (Fig.
2.1B); however this variability in excision gave rise to a chimeric expression pattern of Mcl-1 in
primordial follicle oocytes in neonatal ovaries. Although we still observed a halving in
primordial follicle number shortly after birth (PN7) (Fig. 2.3B), and a doubling in TUNEL
positive apoptotic primordial follicles in PN1 (Fig. 2.3B), the incomplete excision efficiency
prevents us from determining the complete significance of Mcl-1 in neonatal primordial follicle
survival. The use of earlier embryonic Cre Recombinase transgenes Vasa-CREERT2
[26], would
more accurately define that role. Additionally, as concomitant Bax deletion was able to rescue
primordial follicle number in Mcl-1cKO ovaries (Fig. 2.9A), it hints at the interplay between
MCL-1 and BAX as the mediators of primordial follicle oocyte survival. Bax has already
previously been established as the principal pro-apoptotic Bcl-2 core effector member in
regulation of germ cell apoptosis; its interaction with BCL-x determining primordial germ cell
fate [158, 159], and now with MCL-1 regulating primordial follicle fate. Bax deletion resulted in
a marked increase in primordial follicle endowment leading to the prolongation of ovarian
function [156, 157], and has been found to afford protection against a variety of external
environmental inducers of primordial follicle death. Upon activation of follicle growth, Bax-
ablation has been shown to be superfluous in regulation of growing follicle atresia [161],
124
however we have determined that it does lead to a reduction in ovulated oocyte death, even in
Mcl-1-deficient MII oocytes.
Remarkably although the functional interplay between MCL-1 and BAX in oocytes can be
established through genetic studies, our preliminary co-IPs with MCL-1 pulldown did not show
formation of complexes with BAX (Fig. 3.4). This suggests that the interaction between the two
proteins may either be transitory, or instead, due to MCL-1 inhibition of an unknown BH3-only
pro-apoptotic Bcl-2 direct activator of BAX. The current model representing Bcl-2 family
interactions, is a combination of the prior theorized models of interaction [298], and separates
Bcl-2 pro-apoptotic BH3-only members into activators and sensitizers, as described previously.
In this model, sensitizers bind and inhibit the downstream function of anti-apoptotic Bcl-2
members. Anti-apoptotic Bcl-2 members, if not inhibited, can bind and inhibit the action of the
BH3-only activators and the core pro-apoptotic Bcl-2 family effectors. Sensitizers remove the
inhibitory influence of the anti-apoptotic members, allowing the BH3-only activators to stimulate
the effectors, leading to effector intercalation into the mitochondrial membrane, oligomerization,
pore formation and initiation of the apoptotic cascade. Thus MCL-1 mediation of BAX function
in primordial follicle and ovulated oocyte survival may be via MCL-1 inhibition of an activator
of BAX. All three BH3-only activators BIM, BID and PUMA have been revealed to bind BAX
in in vitro studies [131-134] and out of these, BIM and PUMA can bind MCL-1 [136]. PUMA
expression was detected in primordial follicle oocytes of neonatal mice upon γ-irradation, and
deletion of Puma afforded primordial follicle oocytes protection against radiation-induced DNA
damage [176]. BID was demonstrated to be the factor required for BAX activation in regulation
of granulosa cell death [169, 170], however had very little impact on oocytes. Bim expression
was detected in primordial follicle oocytes in rat ovaries, and granulosa cells of mice [173-175],
although Bim ablation resulted in no obvious ovarian phenotype [172], and Bim expression was
125
also determined to be regulated by the transcription factor FOXO3, downstream of the PI3
Kinase pathway [173]. All these factors made BIM a very intriguing option as a BH3-only
activator of BAX; inhibited by MCL-1 in mediation of primordial follicle oocyte fate. We
confirmed MCL-1 and BIM interaction with co-IP’s; however concurrent deletion of Bim in Mcl-
1cKO ovaries, had no apparent rescue phenotype, with histomorphometric analyses revealing
primordial follicle loss comparable to Mcl-1cKO (Fig. 2.9A). To further test whether Bim like
Puma was involved in γ-irradiation-induced activation of primordial follicle cell death, we
irradiated neonatal (PN4) ovaries (0.5-1Gy) and assessed primordial follicle number after 48hrs
(Fig. A1). Bim-ablation resulted in a significant increase in the primordial follicle reserve (Fig.
A1), which is a novel finding; however deletion of Bim did not prevent radiation-induced
primordial follicle loss, but did delay primordial follicle death. Bim-deficient ovaries retained a
significantly higher proportion of dying primordial follicles compared to wildtype controls, 48hrs
after radiation. This implies BIM may possess a subtle yet redundant pro-apoptotic role in
radiation-induced primordial follicle death, with additional pro-apoptotic factors mediating
primordial follicle demise. Conversely, Bim-deletion may instead lead to an impairment in post-
apoptotic primordial follicle clearance. Finally, γ-irradiation did not alter MCL-1 levels or
increase interaction with BIM, as determined by co-IP’s of collected neonatal ovaries (Fig. A2).
Further work will be required to determine the BH3-only activator of BAX in primordial
follicles, possibly via deletion of Bid in primordial follicle oocytes, or a collaborative deletion of
all three activators Bim, Bid, and Puma in case of redundant function.
Intriguingly, although Puma-deletion afforded primordial follicles some protection against γ-
irradiation-induced death, double Puma/Noxa ablation displayed an even greater protection
[176]. Expression of the Bcl-2 pro-apoptotic BH3-only sensitizer Noxa, was noted in primordial
follicle oocytes, but deletion of Noxa alone resulted in no primordial follicle phenotype. NOXA
126
has been established to restrictively bind MCL-1 in vitro [136], and induction of NOXA has also
been attributed with the destabilization of the MCL-1-USP9X de-ubiquitinase complex, leading
to excessive MCL-1 degradation [242]. Future studies should concentrate on the NOXA/MCL-
1/PUMA/BAX axis as a putative mediator of primordial follicle survival.
An additional factor we considered as a putative interacting member of MCL-1 in regulation of
oocyte survival was the pro-apoptotic Bcl-2 effector BOK. BOK protein expression was noted in
fetal human oocytes and oocytes and granulosa of growing follicles [165], which we verified
(data not shown), and perfectly mimics the noted ovarian expression pattern of MCL-1.
Furthermore, BOK was demonstrated to preferentially bind MCL-1 in vitro [164]. However, co-
IP’s performed on total ovarian lysates displayed no interaction of BOK and MCL-1 (Fig. A2),
and total Bok-ablation revealed no direct impact on reproductive function [166]. Further work
will be required to ascertain whether BOK plays any role in regulation of oocyte fate; perhaps
overlapping with other pro-apoptotic Bcl-2 effectors, and hence requiring concomitant deletion
of multiple effectors.
Additionally, as we noted transient nuclear expression of MCL-1 in neonatal primordial follicle
oocytes, we cannot eliminate the possibility that the primordial follicle death associated with
Mcl-1-deficiency may be through another non-apoptosis-related function. MCL-1 has been
identified with roles in cell cycle regulation and DNA repair [272, 273], and thus ablation of
Mcl-1 may result in disruption of either of these outcomes leading to cell defects and eventual
apoptosis. The rescue through Bax-deletion may still be through either prevention of cell death,
via a DNA repair role of Bax, or some additional novel function. Although we establish the
importance of MCL-1 in regulation of primordial follicle survival, further studies are needed to
determine the actual mechanisms. Whether primordial follicle fate is regulated simply by
127
competing levels of MCL-1 vs BAX expression, or whether the presence of any unidentified
activators or sensitizers or additional factors further control the MCL-1/BAX rheostat. In the
next section we will discuss some of the factors we have addressed in our second study that can
regulate primordial follicle survival and growth through regulation of MCL-1.
4.2 Primordial Follicle Growth and Growing Follicle Survival
The granulosa-cell secreted cytokine KL has been well established to modulate primordial
follicle activation, and mediate follicle survival. KL binding to its receptor tyrosine kinase has
been linked to activation of the PI3 Kinase pathway resulting in suppression of FOXO3 activity
[34], in addition to inhibition, via phosphorylation, of GSK-3 [38]. In addition to confirming this
pathway in oocytes using our ex-vivo culture model, we have also verified the additional
downstream GSK-3 inhibitory phosphorylation of MCL-1, previously described in in vitro
studies [238] (Fig. 3.1). The PI3 Kinase pathway has been confirmed to be involved in
primordial follicle activation, i.e. the primordial follicle to primary transition [18, 22, 23, 29, 30,
33], and this further supports the association of this pathway with MCL-1, as we detect an
increased accumulation of MCL-1 with primordial follicle activation by IHC staining (Fig.
2.1A). Recent evidence, leading to impaired KL activation of PI3 Kinase with binding site
mutations, has revealed that KL may not directly mediate primordial follicle activation, but be
required for PI3 Kinase-mediated primordial follicle survival, and primary to secondary growth
[40]. Additional confirmation of the KL-directed elevation of MCL-1 full length expression can
be performed via culture experiments of Mcl-1cKO and control GV oocytes in the presence of
KL. KL treatment should prolong wildtype GV oocyte survival but have no impact on Mcl-1cKO
128
GV’s. However, the possibility remains that PI3 Kinase-mediated MCL-1 activity may be solely
downstream of the KL-activated PI3 Kinase pathway for primordial follicle survival; or a
number of additional growth factors expressed in the primordial to primary transition may
regulate MCL-1 levels.
Further studies will be required to identify these putative growth factors, in addition to a variety
of cytokines and transcription factors that have been verified to modulate Mcl-1 expression in
various in vitro studies [231, 232, 235]. Additionally, if MCL-1 moderation is the defining factor
linked to survival of primordial and the growing follicle pool, molecular mechanisms controlling
MCL-1 stability or translational regulation, must be studied. MCL-1 has an extremely short half-
life [236], and MCL-1 phosphorylation has been linked to recruitment of ubiquitin ligases
MULE, β-TrCP and FBW7 for ubiquitination and proteosomal degradation [238-240].
Counteracting the effects of these ubiquitin ligases are the actions of the de-ubiquitinase USP9X,
which removes ubiquitin tags, thus stabilizing MCL-1 levels [241]. USP9X expression has been
localized to fetal oocytes and oocytes of post-secondary follicles [299], making USP9X a strong
candidate for prolonged MCL-1 stability in growing follicles. Furthermore, USP9X, in genome-
wide association studies have been significantly linked to POF linkage regions on the X-
chromosome [300]. Further studies are required to identify whether USP9X-stabilization of
MCL-1 occurs in the oocyte, and whether its disruption can be linked to POF.
PI3 Kinase mediation of MCL-1 activity through the exogenous granulosa-cell secreted KL
cytokine is one example of granulosa cell management of oocyte survival. Growing follicle cell
death has been well established to occur via a process known as follicle atresia, which is a
granulosa-cell directed means of inducing oocyte death due to the deprivation of support factors
that the granulosa cells provide [146, 147], and is associated with concurrent expression of both
129
apoptotic and autophagic markers [142]. Withdrawal or death of granulosa cells not only
removes a means for secreted cytokine activation of oocyte receptor pathways, but also results in
starvation of the oocyte, as oocyte-granulosa contact is required for the gap junction-mediated
transfer of metabolites and likely ATP to the oocyte [65, 81, 82, 94, 95].
Prior to ovulation, the growing oocyte has been well established to rely on pyruvate breakdown
as the foremost means of energy production [82, 83]; the pyruvate being transferred to the oocyte
by support from the surrounding granulosa cell. We confirmed this reliance with in vitro GV
oocyte cultures (Fig. 3.2A) and also demonstrated that metabolite starvation led to decreased
levels of full length MCL-1 and specifically the mitochondrial matrix-specific MCL-1Matrix
isoform (Fig. 3.2B). Additionally, using IHC, we have shown that sharp downregulation of
MCL-1 accompanies the advent of follicular atresia in post-pubertal ovaries (Fig. 2.1A). To
confidently determine whether MCL-1 downregulation precedes oocyte atresia, a time-dependent
expression analysis can be performed on cultured isolated GV oocytes removed from granulosa
cell support, or cumulus-oocyte-complexes incubated with carbenoxolone [301] a known gap-
junction inhibitor.
The upstream metabolic mediator of MCL-1 levels still remains a mystery. Work by Coloff et al.
have implicated the nutrient-sensitive mTORc1 as one possible upstream factor mediating
translational regulation of MCL-1 levels through activity of the eukaryotic initiation factor
4EBP1 and ribosomal protein rpS6 [255]. Furthermore, AMPK, sensitive to the ATP/AMP
rheostat [200], has been shown to reduce MCL-1 translation through inhibition of mTORc1
activity, upon nutrient starvation [302]. AMPK has been further associated with a role in
maintenance of meiotic arrest and oocyte maturation [303], and the mTORc1 pathway and its
130
downstream components have already been associated with early primordial follicle activation
and survival [18, 22, 33].
Additional studies are required to uncover this possible mechanistic mediation of MCL-1
translation by mTORc1 in oocytes. One possible approach would be the utilization of known
AMPK activators like 5-amino-4-imidazolecarboxamide ribonucleoside (AICAR) [304] or
inhibitors such as compound C [305], on mTOR activity and resultant MCL-1 expression using
WBs in our growing oocyte culture model; in addition to its impact on oocyte survival in our
isolated GV oocyte culture model. Additionally, we can use inhibitors of mTORc1 pathway
activation such as Rapamycin [306] and similarly determine impact on MCL-1 expression and
oocyte survival in our two oocyte culture models. Supplementation or starvation of pyruvate can
be used for effective determination of AMPK pathway inhibition or activation.
Thus we have demonstrated two means of regulation of oocyte MCL-1 levels through granulosa
cell-directed means, cytokine regulation of full length MCL-1, and metabolic regulation of full
length MCL-1 and the MCL-1Matrix
isoform. Previous studies have also demonstrated the dual
nature of control over MCL-1 expression in mouse cell lines. Work by Coloff et al., revealed that
growth factor withdrawal, glucose withdrawal or glycolytic inhibition led to a reduction in MCL-
1 levels, and that restoration of MCL-1 levels using constitutively active AKT, a member of the
PI3 Kinase pathway, required glucose supplementation [255]. Hence we propose that follicular
atresia, instigated upon granulosa cell death or withdrawal, results in a two-pronged reduction of
MCL-1 and the MCL-1Matrix
isoform, via cytokine and metabolic withdrawal; making granulosa
cell-mediated signals the overseers of growing oocyte survival.
131
4.3 Additional Mitochondrial Role for MCL-1 (MCL-1Matrix)
Perciavalle et al. originally identified the 36kDa MCL-1 isoform as a mitochondrial processing
peptidase (MPP) cleaved mitochondrial matrix localized isoform that did not partake in the
traditional anti-apoptotic functions of MCL-1 [221]. They also showed that MCL-1 was required
for proper ATP-Synthase assembly, inferring a role for MCL-1 in functional regulation of
mitochondrial bioenergetics, much akin to the role identified for BCL-x in prevention of leaky
ATP-Synthase activity [220]. Thus, this postulated an additional role of MCL-1 in maintenance
of mitochondrial output, in addition to its anti-apoptotic and anti-autophagic role.
Phenotypic analyses of oocytes obtained from the growing follicle pool have supported this
hypothesis with multiple lines of evidence. In our second study, we observed the putative
interaction of MCL-1 with ATP-Synthase and the ATP Synthase assembly factor (ATPAF)
through co-IP’s in total ovarian lysates (Fig. 3.4). However, total ATP levels were unchanged
between freshly isolated Mcl-1cKO oocytes and controls (Fig. 2.6B). The lack of change in ATP
implied that either Mcl-1cKO oocytes possessed an additional source of exogenous ATP, or the
defective mitochondrial machinery was being overworked to maintain a constant supply of ATP.
It is also possible that our hypothesis was incorrect and MCL-1 was not involved in
mitochondrial bioenergetics. To test the first possibility, we isolated Mcl-1cKO oocytes from the
most obvious source of metabolites, the granulosa cells. Denuded Mcl-1cKO oocytes cultured in
the absence of metabolites died much faster than wildtype controls (Fig. 3.3A). Upon addition of
metabolites, specifically pyruvate or α-ketoglutarate, we noted a significant delay in death of
both Mcl-1cKO oocytes and controls. However this delay in death was not maintained in Mcl-
1cKO oocytes incubated with α-ketoglutarate, to the same extent as control oocytes, as Mcl-
1cKO oocytes still died much faster than wildtype controls in the same treatments. To ascertain
132
the dependence on pyruvate metabolism we cultured these denuded oocytes with inhibitors of
pyruvate uptake by the mitochondria (Fig. 3.3B). Under these parameters, Mcl-1cKO oocytes
still revealed a greater susceptibility to death than wildtype controls. Furthermore, denuded Mcl-
1cKO oocytes matured in vitro failed to create large lipid stores upon maturation as opposed to
wildtype controls (Fig. 3.5B); which indicates that in order to ensure survival, Mcl-1-deficient
oocytes utilize all sources of energy when maintained in the absence of granulosa cell support.
These outcomes actually supported our second postulation as well, that the defective
mitochondrial machinery although disrupted was still functional, and may provide short term
supplies of energy when treated with exogenous metabolites.
Further evidence to support the role of MCL-1, likely the MCL-1Matrix
isoform in mitochondrial
bioenergetics was uncovered in our first study, where we characterized the Mcl-1cKO in growing
and mature oocytes. Mcl-1cKO GV oocytes had increased activation of autophagy (Fig. 2.4B,
C)(Fig. 2.5A, B), which can basally regulate the elimination of malfunctioning organelles, or be
a starvation-induced phenotype resulting in self-digestion [194-196]. The activation of
autophagy may imply that Mcl-1cKO GV oocytes have an excess of damaged organelles (e.g.
mitochondria) and are attempting to rid the cell of them, or that they are self-immolating to
supply the starved oocyte with energy, or perhaps have non-specifically activated autophagy due
to the absence of MCL-1. The latter possibility may be supported by the findings that MCL-1
binds and inhibits Beclin-1, required for autophagosome initiation [204, 219]. However,
Germain et al. have demonstrated that in cortical neurons, the reduction of MCL-1, can lead to
activation of autophagy without simultaneous activation of apoptosis [203]. They propose that
the reduction of MCL-1 precedes activation of either process, dependent on upstream mediators
induced by the source of cellular stress. Thus, in times of starvation, MCL-1 reduction is
regulated by upstream nutrient-sensitive mechanisms, which further stimulates activation of
133
autophagy, but not necessarily apoptosis. Beclin-1 activation has been demonstrated to be reliant
on ULK complex activity and AMPK activity, both initiated by nutrient-deprived conditions
[198, 199, 201, 202].
Hence the activation of autophagy in Mcl-1cKO oocytes is either for provision of additional
energy, or to rid the cell of malfunctioning organelles. Additional evidence for the support of
MCL-1 in mitochondrial bioenergetics was observed using markers of mitochondrial
functionality. Mcl-1cKO oocytes displayed an elevated number of total mitochondria
(MitoTracker Green), but a significant reduction in actively respiring mitochondria (MitoTracker
Red) (Fig. 2.6A). This may imply a compensatory response increasing mitochondria number to
create increased sources of mitochondrial output due to impaired mitochondrial machinery. It
may also imply an increase in mitochondrial fission due to MCL-1 loss or activation of
autophagy. Increases in mitochondrial fission have been suggested as a protective mechanism,
isolating defective mitochondria prior to mitochondrial autophagy (mitophagy) [307, 308].
Determination of the function of autophagic activation in Mcl-1cKO oocytes can be performed
by culture of these GV oocytes and controls in the presence of the autophagy inhibitor 3-methyl
adenine (3-MA) [309]. Dependence of Mcl-1cKO GV oocytes on autophagy for energy supply
should result in advanced death rates if incubated with 3-MA in the absence of pyruvate.
Mcl-1cKO oocytes also displayed an elevation in ROS and specifically mitochondrial derived
superoxides (MitoSOX) (Fig. 2.7A). High ROS levels can result due to defective antioxidant
machinery, or an inhibition/block in the electron transport chain [102-105]. Alternatively,
changes in ROS levels can also be indicative of mitochondrial performance [119, 120], in which
case, high ROS levels imply an overuse of the mitochondrial bioenergetics machinery. Mcl-
1cKO oocytes also exhibited deformed spindle apparatus and increased chromosomal
134
misalignments (Fig. 2.7B), phenotypes that have been associated with high ROS and oxidative
damage, in addition to ATP-depletion in oocytes [114-118]. Conversely, the ATP depletion may
also be causative factor of high oxidative damage of mtDNA [106].
Furthermore, Mcl-1cKO oocytes also displayed a severe reduction in levels of fumarate and
malate, substrates of the TCA cycle (Fig. 2.6B). These various results imply that the absence of
MCL-1 results in mitochondrial malfunction involving faulty mitochondrial bioenergetic output.
To compensate, oocytes appear to activate autophagy, assisted by the lack of MCL-1 inhibition
of Beclin-1, over-employ the existing mitochondrial machinery and may increase mitochondrial
fission, in order to eliminate dysfunctional mitochondria. Further evidence will be required to
validate the increased mitochondrial fission, in addition to confirm the causative factors of
increased ROS; whether it is an overuse of the mitochondrial machinery, or some additional
secondary defect in anti-oxidant mechanisms.
In order to determine whether Mcl-1-deficiency impacts mitochondrial functionality through
defects in mitochondrial fission or fusion, we can use qPCR or WBs on isolated GV oocytes for
changes in expression of markers of either. Regulators of mitochondrial fission include
mitochondrial fission factor (Mff), or dynamin1-like protein (Dnml-1/Drp1); and regulators of
mitochondrial fusion include Mitofusin-1 (Mfn-1) or Mitofusin-2 (Mfn-2).
Concurrent Bax deletion appeared unable to rescue these mitochondrial bioenergetics defects, as
Mcl-1c/BaxDKO oocytes retained elevation of markers of mitochondrial dysfunction.
Furthermore, Mcl-1cKO GV oocytes, despite having an elevation in activated-BAX, did not
display an increase in markers of the apoptotic cascade (Fig. 2.4A). Thus, deletion of Mcl-1 in
oocytes appears to lead to elevated mitochondrial dysfunction, autophagic activation and
decreased metabolic output, but is reinforced by maintenance of granulosa cell support. The fact
135
that these phenotypes are apparently mitochondrial-directed, and seem BAX-independent,
strongly implicates the mitochondrial matrix-restricted MCL-1Matrix
isoform, as the governing
molecule regulating this mitochondrial phenotype, via mediation of ATP-Synthase assembly.
However, we cannot ignore the possibility that BAX plays an additional non-apoptotic role in
growing follicles, perhaps in mitochondrial functionality. BAX has been demonstrated to play a
role in mitochondrial fusion, which is lost upon oligomerization and apoptotic activation [310];
but it remains to be demonstrated whether BAX plays a similar role in growing follicle oocytes.
A complete mitochondrial phenotypic analysis will need to be performed on dual Mcl-1c/BaxKO
GV oocytes and compared to Bax-/-
GVs to determine whether Bax deletion itself contributes to
mitochondrial dysfunction, and whether the Mcl-1c/BaxKO phenotype is more severe. Moreover,
impacts of dual Mcl-1 and Bax deletion will need to be compared to single Mcl-1 or Bax-deleted
GV oocytes to determine expression levels of markers of mitochondrial fusion and fission.
Further confirmation in order to differentiate between the responsibilities of full length MCL-1
and the MCL-1Matrix
isoform may prove to be difficult. We have utilized an inhibitor of MPP, o-
phenanthroline [311] in order to determine how prevention of MCL-1 cleavage may impact
oocyte survival or MCL-1 expression (data not shown), however exaggerated oocyte death tends
to confound our results, as MPP is responsible for post-translational cleavage of a number of
additional mitochondrial factors. MPP, being a metallo-peptidase, has also been found to be
reliant on zinc for effective enzyme activity [312], however confounding results similar to
treatment with o-phenanthroline may be expected. Additional means to isolate cytoplasmic full
length MCL-1 from MCL-1Matrix
isoform functions would be to utilize Mcl-1 expression vector
constructs with mutated MPP cleavage sites for transfection into isolated Mcl-1cKO GV oocytes.
This would prevent formation of the MCL-1Matrix
isoform and allow us to isolate its significant
phenotype. However the zona pellucida membrane which encloses each oocyte has been proven
136
to be impenetrable with common transfection reagents, and as yet the use of carrier proteins to
directly import proteins into the oocyte have also been found to be unsuccessful. Direct
microinjection, may prove to be a solely reliable source of oocyte penetration.
4.4 Meiotic Resumption and Ovulation
In addition to oocyte metabolic support, granulosa cells maintain meiotic arrest through transfer
of cGMP for prevention of cyclic AMP (cAMP) degradation [77-79]. The ovulatory surge of LH
disupts gap junction links and reduces cGMP resulting in meiotic resumption. Follicle atresia can
do the same through withdrawal of granulosa cell contact, in fact the resumption of meiosis is
one of the first steps preceding oocyte atresia [260], and inhibition of meiotic resumption, even
during DNA damage-inducing death stimuli, prevents the activation of apoptotic cellular
fragmentation [261]. However, the initiation of follicle atresia may also be either an apoptosis-
independent event, relying on autophagic and perhaps necrotic cell death, or be mediated by
redundant overlapping efforts of pro-apoptotic Bcl-2 effector proteins, or even other forms of
Bcl-2 independent death [313, 314]. In support of this, GV oocytes cultured in vitro died
predominantly through lysis, which may mimic a joint autophagic/necrotic death, even after GV
breakdown. Only a rare few underwent cellular fragmentation, indicative of apoptotic cell death
[148]. Further markers will be required to identify the form of death in cultured GV oocytes,
which may allow us to determine death molecules regulating follicle atresia as well.
After ovulation, MII oocytes are able to maintain metaphase II arrest for approximately 24hrs,
and then display a proclivity to undergo spontaneous activation and cellular fragmentation,
classified as containing hallmarks of apoptotic cell death [148, 259]. Mcl-1cKO MII oocytes
137
exhibited maintenance of activation of the autophagic pathway (Fig. 2.8A), and also
demonstrated activation of BAX, and increased Caspase activity and Cytochrome c release,
markers of the activated apoptotic cascade (Fig. 2.8B, C). Mcl-1cKO oocytes also revealed an
increased tendency to fragment in culture after 24hrs compared to wildtype controls (Fig. 2.8C),
but intriguingly, Bax deletion was able to completely rescue the Mcl-1cKO fragmentation
phenotype (Fig. 2.10A) and also restored the number of ovulated oocytes to those of wildtype
controls (Fig. 2.9B). We propose that this increased susceptibility to fragment for Mcl-1cKO
oocytes is due to the compromised metabolic phenotype acquired in vivo, resulting in
mitochondrial dysfunction and starvation, contributing to spindle and chromosomal defects.
Oocyte ATP consumption has been revealed to be much higher during phases of oocyte
maturation, both progression through MI and MII [315], and cellular ATP was drastically
reduced upon removal of granulosa cell support. This phenotype resembles that of Mcl-1-ablated
cardiomyocytes, which revealed increases in cell death, mitochondrial dysfunction and impaired
mitochondrial respiration [230]. Concurrent deletion of Bax and Bak, was able to rescue the cell
death, but was unable to rescue the mitochondrial phenotype. Thus, Bax deletion may prevent
fragmentation of these Mcl-1cKO oocytes, but we predict that this will not lead to an improved
breeding performance and restored fertility. Verification of this can be performed by breeding
analyses of the Mcl-1c/BaxDKO females, bred to proven wildtype males. To control for possible
additional phenotypes associated with Bax deletion, we propose a comparative breeding analysis
using stimulated wildtype, Mcl-1cKO, Bax-/-
, and Mcl-1c/BaxDKO females, plugged by proven
wildtype males, and then flushed for embryo transfer into wildtype females for implantation and
delivery.
In this thesis, we have thus identified MCL-1 as an intricate regulator of oocyte survival (Fig.
4.1), by demonstrating its necessity in primordial follicle fate, and the maintenance of the
138
growing oocyte pool through regulation of mitochondrial function and output, in addition to its
anti-apoptotic and anti-autophagic roles. Additionally, we have established two granulosa cell-
directed mechanisms that can mediate MCL-1 oocyte levels through exogenous cytokine and
metabolic support, thus assigning MCL-1 as the primary oocyte survival factor.
In conclusion, we have placed MCL-1 as a key survival factor in the protection of the postnatal
primordial follicle reserve, in addition to maintenance of growing follicle fate. As efficient
reproductive capacity depends on conservation of this primordial follicle pool, and POF remains
a syndrome resulting from its early disruption [3], regulation of MCL-1 and hence follicle
survival, becomes the foremost factor for identifying treatment options. Furthermore, oocyte
quality remains a prime concern in today’s society, as a delay in the child-bearing age has
become more prevalent [274]. Oocyte quality has been demonstrated to deteriorate with
advanced maternal age, resulting in compromised zygotes with increases in aneuploidies,
decreased mitochondrial output and function, oxidative damage and an increased predisposition
to death [4-7, 119, 120, 262]. Our work has revealed that oocyte-specific loss of Mcl-1 results in
an oocyte phenotype with compromised mitochondrial function, increases in ROS, and
chromosomal abnormalities; delegating MCL-1 as an integral component regulating normal
oocyte developmental competence. Additionally, we have proposed a role for MCL-1 in
mediating mitochondrial bioenergetics via the MCL-1Matrix
isoform, giving MCL-1 a direct
means of controlling oocyte quality. Additional work is required to determine whether the in vivo
means of granulosa cell-regulation of oocyte MCL-1 function (cytokine/metabolic), can be
utilized in an exogenous therapeutic approach to improve oocyte quality and reproductive
competence, and preserve ovarian lifespan and function.
139
140
Figure 4.1. Overview of Mechanisms Involved in Oocyte Survival or Death via Regulation
of MCL-1, Presented in this Thesis.
The role of the anti-apoptotic Bcl-2 member Mcl-1 in regulation of primordial germ cell survival cannot be
extrapolated due to Mcl-1 oocyte-specific excision around the time of birth. Thus currently, primordial germ cell
fate has been attributed to the interplay between pro-apoptotic effector BAX and anti-apoptotic BCL-x, during germ
cell migration. In the postnatal ovary, prior to initial recruitment, primordial follicles appear to use MCL-1 to inhibit
a heretofore unverified BH3-only activator of BAX, thus ensuring primordial oocyte survival. Mcl-1 oocyte-
deficiency results in primordial follicle depletion, which is rescued by subsequent Bax deletion; however MCL-1-
BAX interaction does not occur in co-IP’s of neonatal ovaries. MCL-1 may also play additional roles in DNA repair,
which can contribute to primordial follicle death upon Mcl-1ablation.
After initial recruitment, MCL-1 is regulated by two granulosa cell directed mechanisms: extracellular cytokine (via
KL or additional unknown growth factors) activation (blue arrows) of oocyte PI3 Kinase pathway; and gap junction-
mediated metabolic maintenance of MCL-1 levels (green arrows). Levels of MCL-1 full length, and the MCL-1Matrix
isoform have been displayed as reliant on these mechanisms. During follicle atresia, characterized by a withdrawal
of granulosa cell support, and hence these two mechanisms, we verify a reduction in MCL-1 levels via
immunostaining. Additionally, we have observed an increase in oocyte death upon withdrawal of KL cytokine of
metabolites in culture. Follicle atresia is characterized as possessing aspects of both autophagic and apoptotic oocyte
cell death. In Mcl-1cKO GV oocytes, the lack of Mcl-1 leads to activation of autophagy and mitochondrial
dysfunction (impaired mitochondrial bioenergetics), possibly due to disrupted MCL-1Matrix
mediation of ATP
Synthase assembly, leading to decreased mitochondrial output and oocyte starvation. However, maintained
granulosa cell support may prevent an increase in follicle atresia rates, as observed.
Finally, upon ovulation and meiotic resumption, these compromised Mcl-1-deficient oocytes display an increased
susceptibility to death after granulosa cell removal. However this susceptibility can be deterred by simultaneous
ablation of either Bax or pro-apoptotic BH3-only activator Bim. As growing oocytes displayed no direct MCL-1-
BAX interaction through use of co-IP’s, ovulated oocyte survival seems to depends on MCL-1 inhibition of BIM (or
an additional BH3-only member)-activation of BAX.
141
5 REFERENCES
1. Morita, Y. and J.L. Tilly, Oocyte apoptosis: like sand through an hourglass. Dev Biol,
1999. 213(1): p. 1-17.
2. Tilly, J.L., Commuting the death sentence: how oocytes strive to survive. Nat Rev Mol
Cell Biol, 2001. 2(11): p. 838-48.
3. Conway, G.S., Premature ovarian failure. Br Med Bull, 2000. 56(3): p. 643-9.
4. Hunt, P.A. and T.J. Hassold, Human female meiosis: what makes a good egg go bad?
Trends Genet, 2008. 24(2): p. 86-93.
5. Tarin, J.J., S. Perez-Albala, and A. Cano, Consequences on offspring of abnormal
function in ageing gametes. Hum Reprod Update, 2000. 6(6): p. 532-49.
6. Tarin, J.J., S. Perez-Albala, and A. Cano, Cellular and morphological traits of oocytes
retrieved from aging mice after exogenous ovarian stimulation. Biol Reprod, 2001. 65(1):
p. 141-50.
7. Jurisicova, A., et al., Effect of maternal age and conditions of fertilization on
programmed cell death during murine preimplantation embryo development. Mol Hum
Reprod, 1998. 4(2): p. 139-45.
8. Johnson, J., et al., Germline stem cells and follicular renewal in the postnatal mammalian
ovary. Nature, 2004. 428(6979): p. 145-50.
9. Byskov, A.G., Differentiation of mammalian embryonic gonad. Physiol Rev, 1986. 66(1):
p. 71-117.
10. Biason-Lauber, A., WNT4, RSPO1, and FOXL2 in sex development. Semin Reprod Med,
2012. 30(5): p. 387-95.
11. Pepling, M.E. and A.C. Spradling, Female mouse germ cells form synchronously dividing
cysts. Development, 1998. 125(17): p. 3323-8.
12. Pepling, M.E. and A.C. Spradling, Mouse ovarian germ cell cysts undergo programmed
breakdown to form primordial follicles. Dev Biol, 2001. 234(2): p. 339-51.
13. Yao, H.H., The pathway to femaleness: current knowledge on embryonic development of
the ovary. Mol Cell Endocrinol, 2005. 230(1-2): p. 87-93.
14. Hirshfield, A.N., Development of follicles in the mammalian ovary. Int Rev Cytol, 1991.
124: p. 43-101.
15. McLaren, A., Meiosis and differentiation of mouse germ cells. Symp Soc Exp Biol, 1984.
38: p. 7-23.
142
16. McLaren, A., Development of the mammalian gonad: the fate of the supporting cell
lineage. Bioessays, 1991. 13(4): p. 151-6.
17. McLaren, A. and D. Southee, Entry of mouse embryonic germ cells into meiosis. Dev
Biol, 1997. 187(1): p. 107-13.
18. Adhikari, D., et al., Tsc/mTORC1 signaling in oocytes governs the quiescence and
activation of primordial follicles. Hum Mol Genet, 2010. 19(3): p. 397-410.
19. Edson, M.A., A.K. Nagaraja, and M.M. Matzuk, The mammalian ovary from genesis to
revelation. Endocr Rev, 2009. 30(6): p. 624-712.
20. Young, J.M. and A.S. McNeilly, Theca: the forgotten cell of the ovarian follicle.
Reproduction, 2010. 140(4): p. 489-504.
21. McGee, E.A. and A.J. Hsueh, Initial and cyclic recruitment of ovarian follicles. Endocr
Rev, 2000. 21(2): p. 200-14.
22. Castrillon, D.H., et al., Suppression of ovarian follicle activation in mice by the
transcription factor Foxo3a. Science, 2003. 301(5630): p. 215-8.
23. Zheng, W., et al., Functional roles of the phosphatidylinositol 3-kinases (PI3Ks)
signaling in the mammalian ovary. Mol Cell Endocrinol, 2012. 356(1-2): p. 24-30.
24. Hay, N. and N. Sonenberg, Upstream and downstream of mTOR. Genes Dev, 2004.
18(16): p. 1926-45.
25. Lan, Z.J., X. Xu, and A.J. Cooney, Differential oocyte-specific expression of Cre
recombinase activity in GDF-9-iCre, Zp3cre, and Msx2Cre transgenic mice. Biol
Reprod, 2004. 71(5): p. 1469-74.
26. John, G.B., et al., Foxo3 is a PI3K-dependent molecular switch controlling the initiation
of oocyte growth. Dev Biol, 2008. 321(1): p. 197-204.
27. de Vries, W.N., et al., Expression of Cre recombinase in mouse oocytes: a means to study
maternal effect genes. Genesis, 2000. 26(2): p. 110-2.
28. Jagarlamudi, K., et al., Oocyte-specific deletion of Pten in mice reveals a stage-specific
function of PTEN/PI3K signaling in oocytes in controlling follicular activation. PLoS
One, 2009. 4(7): p. e6186.
29. Reddy, P., et al., Oocyte-specific deletion of Pten causes premature activation of the
primordial follicle pool. Science, 2008. 319(5863): p. 611-3.
30. Reddy, P., et al., PDK1 signaling in oocytes controls reproductive aging and lifespan by
manipulating the survival of primordial follicles. Hum Mol Genet, 2009. 18(15): p. 2813-
24.
143
31. Zheng, W., et al., Maternal phosphatidylinositol 3-kinase signalling is crucial for
embryonic genome activation and preimplantation embryogenesis. EMBO Rep, 2010.
11(11): p. 890-5.
32. Brown, C., et al., Subfertility caused by altered follicular development and oocyte growth
in female mice lacking PKB alpha/Akt1. Biol Reprod, 2010. 82(2): p. 246-56.
33. Adhikari, D., et al., Disruption of Tsc2 in oocytes leads to overactivation of the entire
pool of primordial follicles. Mol Hum Reprod, 2009. 15(12): p. 765-70.
34. Reddy, P., et al., Activation of Akt (PKB) and suppression of FKHRL1 in mouse and rat
oocytes by stem cell factor during follicular activation and development. Dev Biol, 2005.
281(2): p. 160-70.
35. Parrott, J.A. and M.K. Skinner, Kit-ligand/stem cell factor induces primordial follicle
development and initiates folliculogenesis. Endocrinology, 1999. 140(9): p. 4262-71.
36. Bedell, M.A., et al., DNA rearrangements located over 100 kb 5' of the Steel (Sl)-coding
region in Steel-panda and Steel-contrasted mice deregulate Sl expression and cause
female sterility by disrupting ovarian follicle development. Genes Dev, 1995. 9(4): p.
455-70.
37. Huang, E.J., et al., The murine steel panda mutation affects kit ligand expression and
growth of early ovarian follicles. Dev Biol, 1993. 157(1): p. 100-9.
38. Liu, L., et al., Phosphorylation and inactivation of glycogen synthase kinase-3 by soluble
kit ligand in mouse oocytes during early follicular development. J Mol Endocrinol, 2007.
38(1-2): p. 137-46.
39. Ali, A., K.P. Hoeflich, and J.R. Woodgett, Glycogen synthase kinase-3: properties,
functions, and regulation. Chem Rev, 2001. 101(8): p. 2527-40.
40. John, G.B., et al., Kit signaling via PI3K promotes ovarian follicle maturation but is
dispensable for primordial follicle activation. Dev Biol, 2009. 331(2): p. 292-9.
41. Layman, L.C. and P.G. McDonough, Mutations of follicle stimulating hormone-beta and
its receptor in human and mouse: genotype/phenotype. Mol Cell Endocrinol, 2000.
161(1-2): p. 9-17.
42. Dierich, A., et al., Impairing follicle-stimulating hormone (FSH) signaling in vivo:
targeted disruption of the FSH receptor leads to aberrant gametogenesis and hormonal
imbalance. Proc Natl Acad Sci U S A, 1998. 95(23): p. 13612-7.
43. Richards, J.S., et al., Novel signaling pathways that control ovarian follicular
development, ovulation, and luteinization. Recent Prog Horm Res, 2002. 57: p. 195-220.
44. Gonzalez-Robayna, I.J., et al., Follicle-Stimulating hormone (FSH) stimulates
phosphorylation and activation of protein kinase B (PKB/Akt) and serum and
144
glucocorticoid-lnduced kinase (Sgk): evidence for A kinase-independent signaling by
FSH in granulosa cells. Mol Endocrinol, 2000. 14(8): p. 1283-300.
45. Liu, Z., et al., FSH and FOXO1 regulate genes in the sterol/steroid and lipid biosynthetic
pathways in granulosa cells. Mol Endocrinol, 2009. 23(5): p. 649-61.
46. Park, Y., et al., Induction of cyclin D2 in rat granulosa cells requires FSH-dependent
relief from FOXO1 repression coupled with positive signals from Smad. J Biol Chem,
2005. 280(10): p. 9135-48.
47. Baker, J., et al., Effects of an Igf1 gene null mutation on mouse reproduction. Mol
Endocrinol, 1996. 10(7): p. 903-18.
48. Zhou, J., et al., Insulin-like growth factor I regulates gonadotropin responsiveness in the
murine ovary. Mol Endocrinol, 1997. 11(13): p. 1924-33.
49. Richards, J.S., et al., Expression of FKHR, FKHRL1, and AFX genes in the rodent ovary:
evidence for regulation by IGF-I, estrogen, and the gonadotropins. Mol Endocrinol,
2002. 16(3): p. 580-99.
50. Cunningham, M.A., Q. Zhu, and J.M. Hammond, FoxO1a can alter cell cycle
progression by regulating the nuclear localization of p27kip in granulosa cells. Mol
Endocrinol, 2004. 18(7): p. 1756-67.
51. Medema, R.H., et al., AFX-like Forkhead transcription factors mediate cell-cycle
regulation by Ras and PKB through p27kip1. Nature, 2000. 404(6779): p. 782-7.
52. Matsuda, F., et al., Expression and function of apoptosis initiator FOXO3 in granulosa
cells during follicular atresia in pig ovaries. J Reprod Dev, 2011. 57(1): p. 151-8.
53. Nakae, J., et al., Insulin regulation of gene expression through the forkhead transcription
factor Foxo1 (Fkhr) requires kinases distinct from Akt. Biochemistry, 2001. 40(39): p.
11768-76.
54. Rizzolio, F., et al., Chromosomal rearrangements in Xq and premature ovarian failure:
mapping of 25 new cases and review of the literature. Hum Reprod, 2006. 21(6): p. 1477-
83.
55. Zinn, A.R. and J.L. Ross, Turner syndrome and haploinsufficiency. Curr Opin Genet
Dev, 1998. 8(3): p. 322-7.
56. Aittomaki, K., et al., Mutation in the follicle-stimulating hormone receptor gene causes
hereditary hypergonadotropic ovarian failure. Cell, 1995. 82(6): p. 959-68.
57. Crisponi, L., et al., FOXL2 inactivation by a translocation 171 kb away: analysis of 500
kb of chromosome 3 for candidate long-range regulatory sequences. Genomics, 2004.
83(5): p. 757-64.
145
58. Kim, S., et al., Epistasis between CYP19A1 and ESR1 polymorphisms is associated with
premature ovarian failure. Fertil Steril, 2011. 95(1): p. 353-6.
59. Laissue, P., et al., Mutations and sequence variants in GDF9 and BMP15 in patients with
premature ovarian failure. Eur J Endocrinol, 2006. 154(5): p. 739-44.
60. Liao, W.X., et al., A new molecular variant of luteinizing hormone associated with
female infertility. Fertil Steril, 1998. 69(1): p. 102-6.
61. Bretherick, K.L., et al., Estrogen receptor alpha gene polymorphisms are associated with
idiopathic premature ovarian failure. Fertil Steril, 2008. 89(2): p. 318-24.
62. Elvin, J.A., et al., Paracrine actions of growth differentiation factor-9 in the mammalian
ovary. Mol Endocrinol, 1999. 13(6): p. 1035-48.
63. Paulini, F. and E.O. Melo, The role of oocyte-secreted factors GDF9 and BMP15 in
follicular development and oogenesis. Reprod Domest Anim, 2011. 46(2): p. 354-61.
64. Eppig, J.J., Oocyte control of ovarian follicular development and function in mammals.
Reproduction, 2001. 122(6): p. 829-38.
65. Su, Y.Q., K. Sugiura, and J.J. Eppig, Mouse oocyte control of granulosa cell development
and function: paracrine regulation of cumulus cell metabolism. Semin Reprod Med,
2009. 27(1): p. 32-42.
66. Anderson, E. and D.F. Albertini, Gap junctions between the oocyte and companion
follicle cells in the mammalian ovary. J Cell Biol, 1976. 71(2): p. 680-6.
67. Ackert, C.L., et al., Intercellular communication via connexin43 gap junctions is
required for ovarian folliculogenesis in the mouse. Dev Biol, 2001. 233(2): p. 258-70.
68. Beyer, E.C., et al., Antisera directed against connexin43 peptides react with a 43-kD
protein localized to gap junctions in myocardium and other tissues. J Cell Biol, 1989.
108(2): p. 595-605.
69. Juneja, S.C., et al., Defects in the germ line and gonads of mice lacking connexin43. Biol
Reprod, 1999. 60(5): p. 1263-70.
70. Simon, A.M., et al., Female infertility in mice lacking connexin 37. Nature, 1997.
385(6616): p. 525-9.
71. Cho, W.K., S. Stern, and J.D. Biggers, Inhibitory effect of dibutyryl cAMP on mouse
oocyte maturation in vitro. J Exp Zool, 1974. 187(3): p. 383-6.
72. Voronina, E. and G.M. Wessel, The regulation of oocyte maturation. Curr Top Dev Biol,
2003. 58: p. 53-110.
73. Horner, K., et al., Rodent oocytes express an active adenylyl cyclase required for meiotic
arrest. Dev Biol, 2003. 258(2): p. 385-96.
146
74. Mehlmann, L.M., T.L. Jones, and L.A. Jaffe, Meiotic arrest in the mouse follicle
maintained by a Gs protein in the oocyte. Science, 2002. 297(5585): p. 1343-5.
75. Webb, R.J., et al., Follicle-stimulating hormone induces a gap junction-dependent
dynamic change in [cAMP] and protein kinase a in mammalian oocytes. Dev Biol, 2002.
246(2): p. 441-54.
76. Bornslaeger, E.A., M.W. Wilde, and R.M. Schultz, Regulation of mouse oocyte
maturation: involvement of cyclic AMP phosphodiesterase and calmodulin. Dev Biol,
1984. 105(2): p. 488-99.
77. Conti, M., et al., Novel signaling mechanisms in the ovary during oocyte maturation and
ovulation. Mol Cell Endocrinol, 2012. 356(1-2): p. 65-73.
78. Norris, R.P., et al., Cyclic GMP from the surrounding somatic cells regulates cyclic AMP
and meiosis in the mouse oocyte. Development, 2009. 136(11): p. 1869-78.
79. Zhang, M. and G. Xia, Hormonal control of mammalian oocyte meiosis at diplotene
stage. Cell Mol Life Sci, 2012. 69(8): p. 1279-88.
80. Vaccari, S., et al., Cyclic GMP signaling is involved in the luteinizing hormone-
dependent meiotic maturation of mouse oocytes. Biol Reprod, 2009. 81(3): p. 595-604.
81. Sutton, M.L., R.B. Gilchrist, and J.G. Thompson, Effects of in-vivo and in-vitro
environments on the metabolism of the cumulus-oocyte complex and its influence on
oocyte developmental capacity. Hum Reprod Update, 2003. 9(1): p. 35-48.
82. Biggers, J.D., D.G. Whittingham, and R.P. Donahue, The pattern of energy metabolism in
the mouse oocyte and zygote. Proc Natl Acad Sci U S A, 1967. 58(2): p. 560-7.
83. Eppig, J.J., Analysis of mouse oogenesis in vitro. Oocyte isolation and the utilization of
exogenous energy sources by growing oocytes. J Exp Zool, 1976. 198(3): p. 375-82.
84. Eppig, J.J., et al., Mouse oocytes regulate metabolic cooperativity between granulosa
cells and oocytes: amino acid transport. Biol Reprod, 2005. 73(2): p. 351-7.
85. Su, Y.Q., et al., Oocyte regulation of metabolic cooperativity between mouse cumulus
cells and oocytes: BMP15 and GDF9 control cholesterol biosynthesis in cumulus cells.
Development, 2008. 135(1): p. 111-21.
86. Sugiura, K., F.L. Pendola, and J.J. Eppig, Oocyte control of metabolic cooperativity
between oocytes and companion granulosa cells: energy metabolism. Dev Biol, 2005.
279(1): p. 20-30.
87. Dumollard, R., et al., Regulation of redox metabolism in the mouse oocyte and embryo.
Development, 2007. 134(3): p. 455-65.
88. Yang, X., et al., Identification of perilipin-2 as a lipid droplet protein regulated in
oocytes during maturation. Reprod Fertil Dev, 2010. 22(8): p. 1262-71.
147
89. Sturmey, R.G., et al., Role of fatty acids in energy provision during oocyte maturation
and early embryo development. Reprod Domest Anim, 2009. 44 Suppl 3: p. 50-8.
90. Dunning, K.R., et al., Beta-oxidation is essential for mouse oocyte developmental
competence and early embryo development. Biol Reprod, 2010. 83(6): p. 909-18.
91. Downs, S.M., P.G. Humpherson, and H.J. Leese, Pyruvate utilization by mouse oocytes is
influenced by meiotic status and the cumulus oophorus. Mol Reprod Dev, 2002. 62(1): p.
113-23.
92. Harris, S.E., et al., Pyruvate and oxygen consumption throughout the growth and
development of murine oocytes. Mol Reprod Dev, 2009. 76(3): p. 231-8.
93. Johnson, M.T., et al., Oxidative metabolism of pyruvate is required for meiotic
maturation of murine oocytes in vivo. Biol Reprod, 2007. 77(1): p. 2-8.
94. Downs, S.M., The influence of glucose, cumulus cells, and metabolic coupling on ATP
levels and meiotic control in the isolated mouse oocyte. Dev Biol, 1995. 167(2): p. 502-
12.
95. Downs, S.M. and A.M. Mastropolo, The participation of energy substrates in the control
of meiotic maturation in murine oocytes. Dev Biol, 1994. 162(1): p. 154-68.
96. Colonna, R. and F. Mangia, Mechanisms of amino acid uptake in cumulus-enclosed
mouse oocytes. Biol Reprod, 1983. 28(4): p. 797-803.
97. Rose-Hellekant, T.A., E.A. Libersky-Williamson, and B.D. Bavister, Energy substrates
and amino acids provided during in vitro maturation of bovine oocytes alter acquisition
of developmental competence. Zygote, 1998. 6(4): p. 285-94.
98. Wycherley, G., M.T. Kane, and A.C. Hynes, Oxidative phosphorylation and the
tricarboxylic acid cycle are essential for normal development of mouse ovarian follicles.
Hum Reprod, 2005. 20(10): p. 2757-63.
99. Nutt, L.K., et al., Metabolic regulation of oocyte cell death through the CaMKII-
mediated phosphorylation of caspase-2. Cell, 2005. 123(1): p. 89-103.
100. Bergeron, L., et al., Defects in regulation of apoptosis in caspase-2-deficient mice. Genes
Dev, 1998. 12(9): p. 1304-14.
101. Sathananthan, A.H. and A.O. Trounson, Mitochondrial morphology during
preimplantational human embryogenesis. Hum Reprod, 2000. 15 Suppl 2: p. 148-59.
102. Boveris, A., E. Cadenas, and A.O. Stoppani, Role of ubiquinone in the mitochondrial
generation of hydrogen peroxide. Biochem J, 1976. 156(2): p. 435-44.
103. Paddenberg, R., et al., Essential role of complex II of the respiratory chain in hypoxia-
induced ROS generation in the pulmonary vasculature. Am J Physiol Lung Cell Mol
Physiol, 2003. 284(5): p. L710-9.
148
104. Turrens, J.F., A. Alexandre, and A.L. Lehninger, Ubisemiquinone is the electron donor
for superoxide formation by complex III of heart mitochondria. Arch Biochem Biophys,
1985. 237(2): p. 408-14.
105. Turrens, J.F. and A. Boveris, Generation of superoxide anion by the NADH
dehydrogenase of bovine heart mitochondria. Biochem J, 1980. 191(2): p. 421-7.
106. Yoneda, M., et al., Oxygen stress induces an apoptotic cell death associated with
fragmentation of mitochondrial genome. Biochem Biophys Res Commun, 1995. 209(2):
p. 723-9.
107. Cree, L.M., et al., A reduction of mitochondrial DNA molecules during embryogenesis
explains the rapid segregation of genotypes. Nat Genet, 2008. 40(2): p. 249-54.
108. Piko, L. and K.D. Taylor, Amounts of mitochondrial DNA and abundance of some
mitochondrial gene transcripts in early mouse embryos. Dev Biol, 1987. 123(2): p. 364-
74.
109. Wai, T., D. Teoli, and E.A. Shoubridge, The mitochondrial DNA genetic bottleneck
results from replication of a subpopulation of genomes. Nat Genet, 2008. 40(12): p.
1484-8.
110. Giles, R.E., et al., Maternal inheritance of human mitochondrial DNA. Proc Natl Acad
Sci U S A, 1980. 77(11): p. 6715-9.
111. Sutovsky, P., et al., Ubiquitinated sperm mitochondria, selective proteolysis, and the
regulation of mitochondrial inheritance in mammalian embryos. Biol Reprod, 2000.
63(2): p. 582-90.
112. El Shourbagy, S.H., et al., Mitochondria directly influence fertilisation outcome in the
pig. Reproduction, 2006. 131(2): p. 233-45.
113. Wai, T., et al., The role of mitochondrial DNA copy number in mammalian fertility. Biol
Reprod, 2010. 83(1): p. 52-62.
114. Zeng, H.T., et al., Low mitochondrial DNA and ATP contents contribute to the absence of
birefringent spindle imaged with PolScope in in vitro matured human oocytes. Hum
Reprod, 2007. 22(6): p. 1681-6.
115. Zeng, H.T., et al., In vitro-matured rat oocytes have low mitochondrial deoxyribonucleic
acid and adenosine triphosphate contents and have abnormal mitochondrial
redistribution. Fertil Steril, 2009. 91(3): p. 900-7.
116. Thouas, G.A., A.O. Trounson, and G.M. Jones, Developmental effects of sublethal
mitochondrial injury in mouse oocytes. Biol Reprod, 2006. 74(5): p. 969-77.
117. Thouas, G.A., et al., Mitochondrial dysfunction in mouse oocytes results in
preimplantation embryo arrest in vitro. Biol Reprod, 2004. 71(6): p. 1936-42.
149
118. Zhang, X., et al., Deficit of mitochondria-derived ATP during oxidative stress impairs
mouse MII oocyte spindles. Cell Res, 2006. 16(10): p. 841-50.
119. Kujjo, L.L., et al., Ceramide and its transport protein (CERT) contribute to deterioration
of mitochondrial structure and function in aging oocytes. Mech Ageing Dev, 2013.
134(1-2): p. 43-52.
120. Kujjo, L.L. and G.I. Perez, Ceramide and mitochondrial function in aging oocytes:
joggling a new hypothesis and old players. Reproduction, 2012. 143(1): p. 1-10.
121. Jurisicova, A. and B.M. Acton, Deadly decisions: the role of genes regulating
programmed cell death in human preimplantation embryo development. Reproduction,
2004. 128(3): p. 281-91.
122. Ziebe, S., et al., Embryo morphology or cleavage stage: how to select the best embryos
for transfer after in-vitro fertilization. Hum Reprod, 1997. 12(7): p. 1545-9.
123. Elmore, S., Apoptosis: a review of programmed cell death. Toxicol Pathol, 2007. 35(4):
p. 495-516.
124. Hsu, S.Y. and A.J. Hsueh, Tissue-specific Bcl-2 protein partners in apoptosis: An
ovarian paradigm. Physiol Rev, 2000. 80(2): p. 593-614.
125. Vaux, D.L., S. Cory, and J.M. Adams, Bcl-2 gene promotes haemopoietic cell survival
and cooperates with c-myc to immortalize pre-B cells. Nature, 1988. 335(6189): p. 440-2.
126. Adams, J.M. and S. Cory, Bcl-2-regulated apoptosis: mechanism and therapeutic
potential. Curr Opin Immunol, 2007. 19(5): p. 488-96.
127. Danial, N.N., BCL-2 family proteins: critical checkpoints of apoptotic cell death. Clin
Cancer Res, 2007. 13(24): p. 7254-63.
128. Youle, R.J. and A. Strasser, The BCL-2 protein family: opposing activities that mediate
cell death. Nat Rev Mol Cell Biol, 2008. 9(1): p. 47-59.
129. Strasser, A., The role of BH3-only proteins in the immune system. Nat Rev Immunol,
2005. 5(3): p. 189-200.
130. Willis, S.N., et al., Proapoptotic Bak is sequestered by Mcl-1 and Bcl-xL, but not Bcl-2,
until displaced by BH3-only proteins. Genes Dev, 2005. 19(11): p. 1294-305.
131. Gallenne, T., et al., Bax activation by the BH3-only protein Puma promotes cell
dependence on antiapoptotic Bcl-2 family members. J Cell Biol, 2009. 185(2): p. 279-90.
132. Harada, H., et al., Survival factor-induced extracellular signal-regulated kinase
phosphorylates BIM, inhibiting its association with BAX and proapoptotic activity. Proc
Natl Acad Sci U S A, 2004. 101(43): p. 15313-7.
150
133. Kim, H., et al., Stepwise activation of BAX and BAK by tBID, BIM, and PUMA initiates
mitochondrial apoptosis. Mol Cell, 2009. 36(3): p. 487-99.
134. Wang, K., et al., BID: a novel BH3 domain-only death agonist. Genes Dev, 1996. 10(22):
p. 2859-69.
135. Ren, D., et al., BID, BIM, and PUMA are essential for activation of the BAX- and BAK-
dependent cell death program. Science, 2010. 330(6009): p. 1390-3.
136. Chen, L., et al., Differential targeting of prosurvival Bcl-2 proteins by their BH3-only
ligands allows complementary apoptotic function. Mol Cell, 2005. 17(3): p. 393-403.
137. Garcia-Saez, A.J., The secrets of the Bcl-2 family. Cell Death Differ, 2012. 19(11): p.
1733-40.
138. Saelens, X., et al., Toxic proteins released from mitochondria in cell death. Oncogene,
2004. 23(16): p. 2861-74.
139. Daugas, E., et al., Mitochondrio-nuclear translocation of AIF in apoptosis and necrosis.
FASEB J, 2000. 14(5): p. 729-39.
140. Norberg, E., S. Orrenius, and B. Zhivotovsky, Mitochondrial regulation of cell death:
processing of apoptosis-inducing factor (AIF). Biochem Biophys Res Commun, 2010.
396(1): p. 95-100.
141. Li, L.Y., X. Luo, and X. Wang, Endonuclease G is an apoptotic DNase when released
from mitochondria. Nature, 2001. 412(6842): p. 95-9.
142. Escobar, M.L., et al., Combined apoptosis and autophagy, the process that eliminates the
oocytes of atretic follicles in immature rats. Apoptosis, 2008. 13(10): p. 1253-66.
143. Lobascio, A.M., et al., Analysis of programmed cell death in mouse fetal oocytes.
Reproduction, 2007. 134(2): p. 241-52.
144. De Felici, M., A.M. Lobascio, and F.G. Klinger, Cell death in fetal oocytes: many
players for multiple pathways. Autophagy, 2008. 4(2): p. 240-2.
145. Tingen, C.M., et al., Prepubertal primordial follicle loss in mice is not due to classical
apoptotic pathways. Biol Reprod, 2009. 81(1): p. 16-25.
146. Byskov, A.G., Cell kinetic studies of follicular atresia in the mouse ovary. J Reprod
Fertil, 1974. 37(2): p. 277-85.
147. Devine, P.J., et al., Ultrastructural evaluation of oocytes during atresia in rat ovarian
follicles. Biol Reprod, 2000. 63(5): p. 1245-52.
148. Perez, G.I., X.J. Tao, and J.L. Tilly, Fragmentation and death (a.k.a. apoptosis) of
ovulated oocytes. Mol Hum Reprod, 1999. 5(5): p. 414-20.
151
149. Tilly, J.L., et al., Expression of members of the bcl-2 gene family in the immature rat
ovary: equine chorionic gonadotropin-mediated inhibition of granulosa cell apoptosis is
associated with decreased bax and constitutive bcl-2 and bcl-xlong messenger
ribonucleic acid levels. Endocrinology, 1995. 136(1): p. 232-41.
150. Knudson, C.M., et al., Bax-deficient mice with lymphoid hyperplasia and male germ cell
death. Science, 1995. 270(5233): p. 96-9.
151. Jurisicova, A., et al., Expression and regulation of genes associated with cell death
during murine preimplantation embryo development. Mol Reprod Dev, 1998. 51(3): p.
243-53.
152. Takai, Y., et al., Bax, caspase-2, and caspase-3 are required for ovarian follicle loss
caused by 4-vinylcyclohexene diepoxide exposure of female mice in vivo. Endocrinology,
2003. 144(1): p. 69-74.
153. Kujjo, L.L., et al., Enhancing survival of mouse oocytes following chemotherapy or aging
by targeting Bax and Rad51. PLoS One, 2010. 5(2): p. e9204.
154. Perez, G.I., et al., Apoptosis-associated signaling pathways are required for
chemotherapy-mediated female germ cell destruction. Nat Med, 1997. 3(11): p. 1228-32.
155. Matikainen, T., et al., Aromatic hydrocarbon receptor-driven Bax gene expression is
required for premature ovarian failure caused by biohazardous environmental chemicals.
Nat Genet, 2001. 28(4): p. 355-60.
156. Perez, G.I., et al., Absence of the proapoptotic Bax protein extends fertility and alleviates
age-related health complications in female mice. Proc Natl Acad Sci U S A, 2007.
104(12): p. 5229-34.
157. Perez, G.I., et al., Prolongation of ovarian lifespan into advanced chronological age by
Bax-deficiency. Nat Genet, 1999. 21(2): p. 200-3.
158. Rucker, E.B., 3rd, et al., Bcl-x and Bax regulate mouse primordial germ cell survival and
apoptosis during embryogenesis. Mol Endocrinol, 2000. 14(7): p. 1038-52.
159. Stallock, J., et al., The pro-apoptotic gene Bax is required for the death of ectopic
primordial germ cells during their migration in the mouse embryo. Development, 2003.
130(26): p. 6589-97.
160. Greenfeld, C.R., et al., BAX regulates follicular endowment in mice. Reproduction, 2007.
133(5): p. 865-76.
161. Greenfeld, C.R., et al., BAX is involved in regulating follicular growth, but is dispensable
for follicle atresia in adult mouse ovaries. Reproduction, 2007. 133(1): p. 107-16.
162. Krajewski, S., M. Krajewska, and J.C. Reed, Immuno-histochemical analysis of in vivo
patterns of Bak expression, a proapoptotic member of the Bcl-2 protein family. Cancer
Res, 1996. 56(12): p. 2849-55.
152
163. Lindsten, T., et al., The combined functions of proapoptotic Bcl-2 family members bak
and bax are essential for normal development of multiple tissues. Mol Cell, 2000. 6(6): p.
1389-99.
164. Hsu, S.Y., et al., Bok is a pro-apoptotic Bcl-2 protein with restricted expression in
reproductive tissues and heterodimerizes with selective anti-apoptotic Bcl-2 family
members. Proc Natl Acad Sci U S A, 1997. 94(23): p. 12401-6.
165. Jaaskelainen, M., et al., Regulation of cell death in human fetal and adult ovaries--role of
Bok and Bcl-X(L). Mol Cell Endocrinol, 2010. 330(1-2): p. 17-24.
166. Ke, F., et al., BCL-2 family member BOK is widely expressed but its loss has only
minimal impact in mice. Cell Death Differ, 2012. 19(6): p. 915-25.
167. Kaipia, A., S.Y. Hsu, and A.J. Hsueh, Expression and function of a proapoptotic Bcl-2
family member Bcl-XL/Bcl-2-associated death promoter (BAD) in rat ovary.
Endocrinology, 1997. 138(12): p. 5497-504.
168. Jurisicova, A., et al., Maternal exposure to polycyclic aromatic hydrocarbons diminishes
murine ovarian reserve via induction of Harakiri. J Clin Invest, 2007. 117(12): p. 3971-8.
169. Sai, T., et al., Bid and Bax are involved in granulosa cell apoptosis during follicular
atresia in porcine ovaries. J Reprod Dev, 2011. 57(3): p. 421-7.
170. Sai, T., et al., Effect of RNA interference of BID and BAX mRNAs on apoptosis in
granulosa cell-derived KGN cells. J Reprod Dev, 2012. 58(1): p. 112-6.
171. Hsu, S.Y., P. Lin, and A.J. Hsueh, BOD (Bcl-2-related ovarian death gene) is an ovarian
BH3 domain-containing proapoptotic Bcl-2 protein capable of dimerization with diverse
antiapoptotic Bcl-2 members. Mol Endocrinol, 1998. 12(9): p. 1432-40.
172. Bouillet, P., et al., Proapoptotic Bcl-2 relative Bim required for certain apoptotic
responses, leukocyte homeostasis, and to preclude autoimmunity. Science, 1999.
286(5445): p. 1735-8.
173. Liu, H., et al., FOXO3a is involved in the apoptosis of naked oocytes and oocytes of
primordial follicles from neonatal rat ovaries. Biochem Biophys Res Commun, 2009.
381(4): p. 722-7.
174. O'Reilly, L.A., et al., The proapoptotic BH3-only protein bim is expressed in
hematopoietic, epithelial, neuronal, and germ cells. Am J Pathol, 2000. 157(2): p. 449-
61.
175. Wang, X.L., et al., Follicle-stimulating hormone regulates pro-apoptotic protein Bcl-2-
interacting mediator of cell death-extra long (BimEL)-induced porcine granulosa cell
apoptosis. J Biol Chem, 2012. 287(13): p. 10166-77.
153
176. Kerr, J.B., et al., DNA damage-induced primordial follicle oocyte apoptosis and loss of
fertility require TAp63-mediated induction of Puma and Noxa. Mol Cell, 2012. 48(3): p.
343-52.
177. Livera, G., et al., p63 null mutation protects mouse oocytes from radio-induced
apoptosis. Reproduction, 2008. 135(1): p. 3-12.
178. Suh, E.K., et al., p63 protects the female germ line during meiotic arrest. Nature, 2006.
444(7119): p. 624-8.
179. Ratts, V.S., et al., Ablation of bcl-2 gene expression decreases the numbers of oocytes
and primordial follicles established in the post-natal female mouse gonad.
Endocrinology, 1995. 136(8): p. 3665-8.
180. Flaws, J.A., et al., Effect of bcl-2 on the primordial follicle endowment in the mouse
ovary. Biol Reprod, 2001. 64(4): p. 1153-9.
181. Jones, R.L. and M.E. Pepling, Role of the antiapoptotic proteins BCL2 and MCL1 in the
neonatal mouse ovary. Biol Reprod, 2013. 88(2): p. 46.
182. Gursoy, E., et al., Expression and localisation of Bcl-2 and Bax proteins in developing
rat ovary. Res Vet Sci, 2008. 84(1): p. 56-61.
183. Hsu, S.Y., et al., Targeted overexpression of Bcl-2 in ovaries of transgenic mice leads to
decreased follicle apoptosis, enhanced folliculogenesis, and increased germ cell
tumorigenesis. Endocrinology, 1996. 137(11): p. 4837-43.
184. Krajewski, S., et al., Immuno-histochemical analysis of in vivo patterns of Bcl-X
expression. Cancer Res, 1994. 54(21): p. 5501-7.
185. Motoyama, N., et al., Massive cell death of immature hematopoietic cells and neurons in
Bcl-x-deficient mice. Science, 1995. 267(5203): p. 1506-10.
186. Johnson, A.L., J.T. Bridgham, and T. Jensen, Bcl-X(LONG) protein expression and
phosphorylation in granulosa cells. Endocrinology, 1999. 140(10): p. 4521-9.
187. Riedlinger, G., et al., Bcl-x is not required for maintenance of follicles and corpus luteum
in the postnatal mouse ovary. Biol Reprod, 2002. 66(2): p. 438-44.
188. Inohara, N., et al., Diva, a Bcl-2 homologue that binds directly to Apaf-1 and induces
BH3-independent cell death. J Biol Chem, 1998. 273(49): p. 32479-86.
189. Guillemin, Y., et al., Oocytes and early embryos selectively express the survival factor
BCL2L10. J Mol Med (Berl), 2009. 87(9): p. 923-40.
190. Russell, H.R., et al., Murine ovarian development is not affected by inactivation of the
bcl-2 family member diva. Mol Cell Biol, 2002. 22(19): p. 6866-70.
154
191. Print, C.G., et al., Apoptosis regulator bcl-w is essential for spermatogenesis but appears
otherwise redundant. Proc Natl Acad Sci U S A, 1998. 95(21): p. 12424-31.
192. Hartley, P.S., et al., Developmental changes in expression of myeloid cell leukemia-1 in
human germ cells during oogenesis and early folliculogenesis. J Clin Endocrinol Metab,
2002. 87(7): p. 3417-27.
193. Leo, C.P., et al., Characterization of the antiapoptotic Bcl-2 family member myeloid cell
leukemia-1 (Mcl-1) and the stimulation of its message by gonadotropins in the rat ovary.
Endocrinology, 1999. 140(12): p. 5469-77.
194. Edinger, A.L. and C.B. Thompson, Death by design: apoptosis, necrosis and autophagy.
Curr Opin Cell Biol, 2004. 16(6): p. 663-9.
195. Eisenberg-Lerner, A., et al., Life and death partners: apoptosis, autophagy and the cross-
talk between them. Cell Death Differ, 2009. 16(7): p. 966-75.
196. Kroemer, G., G. Marino, and B. Levine, Autophagy and the integrated stress response.
Mol Cell, 2010. 40(2): p. 280-93.
197. Noda, T. and Y. Ohsumi, Tor, a phosphatidylinositol kinase homologue, controls
autophagy in yeast. J Biol Chem, 1998. 273(7): p. 3963-6.
198. Mizushima, N., The role of the Atg1/ULK1 complex in autophagy regulation. Curr Opin
Cell Biol, 2010. 22(2): p. 132-9.
199. Russell, R.C., et al., ULK1 induces autophagy by phosphorylating Beclin-1 and
activating VPS34 lipid kinase. Nat Cell Biol, 2013.
200. Hardie, D.G., AMP-activated protein kinase as a drug target. Annu Rev Pharmacol
Toxicol, 2007. 47: p. 185-210.
201. Kim, J., et al., Differential regulation of distinct Vps34 complexes by AMPK in nutrient
stress and autophagy. Cell, 2013. 152(1-2): p. 290-303.
202. Egan, D.F., et al., Phosphorylation of ULK1 (hATG1) by AMP-activated protein kinase
connects energy sensing to mitophagy. Science, 2011. 331(6016): p. 456-61.
203. Germain, M., et al., MCL-1 is a stress sensor that regulates autophagy in a
developmentally regulated manner. EMBO J, 2011. 30(2): p. 395-407.
204. Maiuri, M.C., et al., Functional and physical interaction between Bcl-X(L) and a BH3-
like domain in Beclin-1. EMBO J, 2007. 26(10): p. 2527-39.
205. Pattingre, S., et al., Bcl-2 antiapoptotic proteins inhibit Beclin 1-dependent autophagy.
Cell, 2005. 122(6): p. 927-39.
206. Hailey, D.W., et al., Mitochondria supply membranes for autophagosome biogenesis
during starvation. Cell, 2010. 141(4): p. 656-67.
155
207. Fujita, N., et al., The Atg16L complex specifies the site of LC3 lipidation for membrane
biogenesis in autophagy. Mol Biol Cell, 2008. 19(5): p. 2092-100.
208. Johansen, T. and T. Lamark, Selective autophagy mediated by autophagic adapter
proteins. Autophagy, 2011. 7(3): p. 279-96.
209. Morita, E. and T. Yoshimori, Membrane recruitment of autophagy proteins in selective
autophagy. Hepatol Res, 2012. 42(5): p. 435-41.
210. Sou, Y.S., et al., The Atg8 conjugation system is indispensable for proper development of
autophagic isolation membranes in mice. Mol Biol Cell, 2008. 19(11): p. 4762-75.
211. Huynh, K.K., et al., LAMP proteins are required for fusion of lysosomes with
phagosomes. EMBO J, 2007. 26(2): p. 313-24.
212. Chen, Y. and L. Yu, Autophagic lysosome reformation. Exp Cell Res, 2013. 319(2): p.
142-6.
213. Tsukamoto, S., A. Kuma, and N. Mizushima, The role of autophagy during the oocyte-to-
embryo transition. Autophagy, 2008. 4(8): p. 1076-8.
214. Escobar Sanchez, M.L., O.M. Echeverria Martinez, and G.H. Vazquez-Nin, Immuno-
histochemical and ultrastructural visualization of different routes of oocyte elimination in
adult rats. Eur J Histochem, 2012. 56(2): p. e17.
215. Gaytan, M., et al., Immunolocalization of beclin 1, a bcl-2-binding, autophagy-related
protein, in the human ovary: possible relation to life span of corpus luteum. Cell Tissue
Res, 2008. 331(2): p. 509-17.
216. Gawriluk, T.R., et al., Autophagy is a cell survival program for female germ cells in the
murine ovary. Reproduction, 2011. 141(6): p. 759-65.
217. Andersen, J.L. and S. Kornbluth, The tangled circuitry of metabolism and apoptosis. Mol
Cell, 2013. 49(3): p. 399-410.
218. Liang, X.H., et al., Protection against fatal Sindbis virus encephalitis by beclin, a novel
Bcl-2-interacting protein. J Virol, 1998. 72(11): p. 8586-96.
219. Erlich, S., et al., Differential interactions between Beclin 1 and Bcl-2 family members.
Autophagy, 2007. 3(6): p. 561-8.
220. Alavian, K.N., et al., Bcl-xL regulates metabolic efficiency of neurons through
interaction with the mitochondrial F1FO ATP synthase. Nat Cell Biol, 2011. 13(10): p.
1224-33.
221. Perciavalle, R.M., et al., Anti-apoptotic MCL-1 localizes to the mitochondrial matrix and
couples mitochondrial fusion to respiration. Nat Cell Biol, 2012. 14(6): p. 575-83.
156
222. Maurer, U., et al., Glycogen synthase kinase-3 regulates mitochondrial outer membrane
permeabilization and apoptosis by destabilization of MCL-1. Mol Cell, 2006. 21(6): p.
749-60.
223. Zhao, Y., et al., Glycogen synthase kinase 3alpha and 3beta mediate a glucose-sensitive
antiapoptotic signaling pathway to stabilize Mcl-1. Mol Cell Biol, 2007. 27(12): p. 4328-
39.
224. Kozopas, K.M., et al., MCL1, a gene expressed in programmed myeloid cell
differentiation, has sequence similarity to BCL2. Proc Natl Acad Sci U S A, 1993. 90(8):
p. 3516-20.
225. Rinkenberger, J.L., et al., Mcl-1 deficiency results in peri-implantation embryonic
lethality. Genes Dev, 2000. 14(1): p. 23-7.
226. Opferman, J.T., et al., Development and maintenance of B and T lymphocytes requires
antiapoptotic MCL-1. Nature, 2003. 426(6967): p. 671-6.
227. Opferman, J.T., et al., Obligate role of anti-apoptotic MCL-1 in the survival of
hematopoietic stem cells. Science, 2005. 307(5712): p. 1101-4.
228. Dzhagalov, I., A. St John, and Y.W. He, The antiapoptotic protein Mcl-1 is essential for
the survival of neutrophils but not macrophages. Blood, 2007. 109(4): p. 1620-6.
229. Steimer, D.A., et al., Selective roles for antiapoptotic MCL-1 during granulocyte
development and macrophage effector function. Blood, 2009. 113(12): p. 2805-15.
230. Wang, X., et al., Deletion of MCL-1 causes lethal cardiac failure and mitochondrial
dysfunction. Genes Dev, 2013. 27(12): p. 1351-64.
231. Thomas, L.W., C. Lam, and S.W. Edwards, Mcl-1; the molecular regulation of protein
function. FEBS Lett, 2010. 584(14): p. 2981-9.
232. Gores, G.J. and S.H. Kaufmann, Selectively targeting Mcl-1 for the treatment of acute
myelogenous leukemia and solid tumors. Genes Dev, 2012. 26(4): p. 305-11.
233. Bae, J., et al., MCL-1S, a splicing variant of the antiapoptotic BCL-2 family member
MCL-1, encodes a proapoptotic protein possessing only the BH3 domain. J Biol Chem,
2000. 275(33): p. 25255-61.
234. Kim, J.H., et al., MCL-1ES, a novel variant of MCL-1, associates with MCL-1L and
induces mitochondrial cell death. FEBS Lett, 2009. 583(17): p. 2758-64.
235. Huang, H.M., C.J. Huang, and J.J. Yen, Mcl-1 is a common target of stem cell factor and
interleukin-5 for apoptosis prevention activity via MEK/MAPK and PI-3K/Akt pathways.
Blood, 2000. 96(5): p. 1764-71.
236. Adams, K.W. and G.M. Cooper, Rapid turnover of mcl-1 couples translation to cell
survival and apoptosis. J Biol Chem, 2007. 282(9): p. 6192-200.
157
237. Nijhawan, D., et al., Elimination of Mcl-1 is required for the initiation of apoptosis
following ultraviolet irradiation. Genes Dev, 2003. 17(12): p. 1475-86.
238. Ding, Q., et al., Degradation of Mcl-1 by beta-TrCP mediates glycogen synthase kinase
3-induced tumor suppression and chemosensitization. Mol Cell Biol, 2007. 27(11): p.
4006-17.
239. Wertz, I.E., et al., Sensitivity to antitubulin chemotherapeutics is regulated by MCL1 and
FBW7. Nature, 2011. 471(7336): p. 110-4.
240. Zhong, Q., et al., Mule/ARF-BP1, a BH3-only E3 ubiquitin ligase, catalyzes the
polyubiquitination of Mcl-1 and regulates apoptosis. Cell, 2005. 121(7): p. 1085-95.
241. Schwickart, M., et al., Deubiquitinase USP9X stabilizes MCL1 and promotes tumour cell
survival. Nature, 2010. 463(7277): p. 103-7.
242. Gomez-Bougie, P., et al., Noxa controls Mule-dependent Mcl-1 ubiquitination through
the regulation of the Mcl-1/USP9X interaction. Biochem Biophys Res Commun, 2011.
413(3): p. 460-4.
243. Clohessy, J.G., J. Zhuang, and H.J. Brady, Characterisation of Mcl-1 cleavage during
apoptosis of haematopoietic cells. Br J Haematol, 2004. 125(5): p. 655-65.
244. Weng, C., et al., Specific cleavage of Mcl-1 by caspase-3 in tumor necrosis factor-related
apoptosis-inducing ligand (TRAIL)-induced apoptosis in Jurkat leukemia T cells. J Biol
Chem, 2005. 280(11): p. 10491-500.
245. Herrant, M., et al., Cleavage of Mcl-1 by caspases impaired its ability to counteract Bim-
induced apoptosis. Oncogene, 2004. 23(47): p. 7863-73.
246. Han, J., et al., Degradation of Mcl-1 by granzyme B: implications for Bim-mediated
mitochondrial apoptotic events. J Biol Chem, 2004. 279(21): p. 22020-9.
247. Han, J., et al., Disruption of Mcl-1.Bim complex in granzyme B-mediated mitochondrial
apoptosis. J Biol Chem, 2005. 280(16): p. 16383-92.
248. Jamil, S., et al., A proteolytic fragment of Mcl-1 exhibits nuclear localization and
regulates cell growth by interaction with Cdk1. Biochem J, 2005. 387(Pt 3): p. 659-67.
249. Kobayashi, S., et al., Serine 64 phosphorylation enhances the antiapoptotic function of
Mcl-1. J Biol Chem, 2007. 282(25): p. 18407-17.
250. Domina, A.M., et al., MCL1 is phosphorylated in the PEST region and stabilized upon
ERK activation in viable cells, and at additional sites with cytotoxic okadaic acid or
taxol. Oncogene, 2004. 23(31): p. 5301-15.
251. Ding, Q., et al., Down-regulation of myeloid cell leukemia-1 through inhibiting Erk/Pin 1
pathway by sorafenib facilitates chemosensitization in breast cancer. Cancer Res, 2008.
68(15): p. 6109-17.
158
252. Inoshita, S., et al., Phosphorylation and inactivation of myeloid cell leukemia 1 by JNK in
response to oxidative stress. J Biol Chem, 2002. 277(46): p. 43730-4.
253. Kodama, Y., et al., Antiapoptotic effect of c-Jun N-terminal Kinase-1 through Mcl-1
stabilization in TNF-induced hepatocyte apoptosis. Gastroenterology, 2009. 136(4): p.
1423-34.
254. Morel, C., et al., Mcl-1 integrates the opposing actions of signaling pathways that
mediate survival and apoptosis. Mol Cell Biol, 2009. 29(14): p. 3845-52.
255. Coloff, J.L., et al., Akt-dependent glucose metabolism promotes Mcl-1 synthesis to
maintain cell survival and resistance to Bcl-2 inhibition. Cancer Res, 2011. 71(15): p.
5204-13.
256. Huang, C.R. and H.F. Yang-Yen, The fast-mobility isoform of mouse Mcl-1 is a
mitochondrial matrix-localized protein with attenuated anti-apoptotic activity. FEBS
Lett, 2010. 584(15): p. 3323-30.
257. Boya, P., et al., Inhibition of macroautophagy triggers apoptosis. Mol Cell Biol, 2005.
25(3): p. 1025-40.
258. Wirawan, E., et al., Caspase-mediated cleavage of Beclin-1 inactivates Beclin-1-induced
autophagy and enhances apoptosis by promoting the release of proapoptotic factors from
mitochondria. Cell Death Dis, 2010. 1: p. e18.
259. Takase, K., M. Ishikawa, and H. Hoshiai, Apoptosis in the degeneration process of
unfertilized mouse ova. Tohoku J Exp Med, 1995. 175(1): p. 69-76.
260. Lefevre, B., et al., In vivo changes in oocyte germinal vesicle related to follicular quality
and size at mid-follicular phase during stimulated cycles in the cynomolgus monkey.
Reprod Nutr Dev, 1989. 29(5): p. 523-31.
261. Jurisicova, A., et al., Molecular requirements for doxorubicin-mediated death in murine
oocytes. Cell Death Differ, 2006. 13(9): p. 1466-74.
262. Hamatani, T., et al., Age-associated alteration of gene expression patterns in mouse
oocytes. Hum Mol Genet, 2004. 13(19): p. 2263-78.
263. Lewandoski, M., K.M. Wassarman, and G.R. Martin, Zp3-cre, a transgenic mouse line
for the activation or inactivation of loxP-flanked target genes specifically in the female
germ line. Curr Biol, 1997. 7(2): p. 148-51.
264. Lobe, C.G., et al., Z/AP, a double reporter for cre-mediated recombination. Dev Biol,
1999. 208(2): p. 281-92.
265. Madisen, L., et al., A robust and high-throughput Cre reporting and characterization
system for the whole mouse brain. Nat Neurosci, 2010. 13(1): p. 133-40.
159
266. Takeuchi, O., et al., Essential role of BAX,BAK in B cell homeostasis and prevention of
autoimmune disease. Proc Natl Acad Sci U S A, 2005. 102(32): p. 11272-7.
267. Chance, B., et al., Oxidation-reduction ratio studies of mitochondria in freeze-trapped
samples. NADH and flavoprotein fluorescence signals. J Biol Chem, 1979. 254(11): p.
4764-71.
268. Eng, J., R.M. Lynch, and R.S. Balaban, Nicotinamide adenine dinucleotide fluorescence
spectroscopy and imaging of isolated cardiac myocytes. Biophys J, 1989. 55(4): p. 621-
30.
269. Reinert, K.C., et al., Flavoprotein autofluorescence imaging in the cerebellar cortex in
vivo. J Neurosci Res, 2007. 85(15): p. 3221-32.
270. Chi, M.M., A. Hoehn, and K.H. Moley, Metabolic changes in the glucose-induced
apoptotic blastocyst suggest alterations in mitochondrial physiology. Am J Physiol
Endocrinol Metab, 2002. 283(2): p. E226-32.
271. Kang, R., et al., The Beclin 1 network regulates autophagy and apoptosis. Cell Death
Differ, 2011. 18(4): p. 571-80.
272. Arbour, N., et al., Mcl-1 is a key regulator of apoptosis during CNS development and
after DNA damage. J Neurosci, 2008. 28(24): p. 6068-78.
273. Fujise, K., et al., Regulation of apoptosis and cell cycle progression by MCL1.
Differential role of proliferating cell nuclear antigen. J Biol Chem, 2000. 275(50): p.
39458-65.
274. Matthews, T.J. and B.E. Hamilton, Delayed childbearing: more women are having their
first child later in life. NCHS Data Brief, 2009(21): p. 1-8.
275. Trivigno, D., et al., Deubiquitinase USP9x confers radioresistance through stabilization
of Mcl-1. Neoplasia, 2012. 14(10): p. 893-904.
276. Embi, N., D.B. Rylatt, and P. Cohen, Glycogen synthase kinase-3 from rabbit skeletal
muscle. Separation from cyclic-AMP-dependent protein kinase and phosphorylase
kinase. Eur J Biochem, 1980. 107(2): p. 519-27.
277. Welsh, G.I., C. Wilson, and C.G. Proud, GSK3: a SHAGGY frog story. Trends Cell Biol,
1996. 6(7): p. 274-9.
278. Lochhead, P.A., et al., Inhibition of GSK-3 selectively reduces glucose-6-phosphatase
and phosphatase and phosphoenolypyruvate carboxykinase gene expression. Diabetes,
2001. 50(5): p. 937-46.
279. Liberman, Z. and H. Eldar-Finkelman, Serine 332 phosphorylation of insulin receptor
substrate-1 by glycogen synthase kinase-3 attenuates insulin signaling. J Biol Chem,
2005. 280(6): p. 4422-8.
160
280. Leese, H.J. and A.M. Barton, Pyruvate and glucose uptake by mouse ova and
preimplantation embryos. J Reprod Fertil, 1984. 72(1): p. 9-13.
281. Wensveen, F.M., et al., Apoptosis induced by overall metabolic stress converges on the
Bcl-2 family proteins Noxa and Mcl-1. Apoptosis, 2011. 16(7): p. 708-21.
282. Halestrap, A.P., The mitochondrial pyruvate carrier. Kinetics and specificity for
substrates and inhibitors. Biochem J, 1975. 148(1): p. 85-96.
283. Johnson, I.D., H.C. Kang, and R.P. Haugland, Fluorescent membrane probes
incorporating dipyrrometheneboron difluoride fluorophores. Anal Biochem, 1991.
198(2): p. 228-37.
284. Luo, S. and D.C. Rubinsztein, BCL2L11/BIM: a novel molecular link between autophagy
and apoptosis. Autophagy, 2013. 9(1): p. 104-5.
285. Mortimore, G.E. and A.R. Poso, Intracellular protein catabolism and its control during
nutrient deprivation and supply. Annu Rev Nutr, 1987. 7: p. 539-64.
286. Downs, S.M., J.L. Mosey, and J. Klinger, Fatty acid oxidation and meiotic resumption in
mouse oocytes. Mol Reprod Dev, 2009. 76(9): p. 844-53.
287. Faddy, M.J., et al., Accelerated disappearance of ovarian follicles in mid-life:
implications for forecasting menopause. Hum Reprod, 1992. 7(10): p. 1342-6.
288. Richardson, S.J., V. Senikas, and J.F. Nelson, Follicular depletion during the
menopausal transition: evidence for accelerated loss and ultimate exhaustion. J Clin
Endocrinol Metab, 1987. 65(6): p. 1231-7.
289. Hassold, T. and P. Hunt, Maternal age and chromosomally abnormal pregnancies: what
we know and what we wish we knew. Curr Opin Pediatr, 2009. 21(6): p. 703-8.
290. Titus, S., et al., Impairment of BRCA1-related DNA double-strand break repair leads to
ovarian aging in mice and humans. Sci Transl Med, 2013. 5(172): p. 172ra21.
291. Andersen, C.Y., et al., Cryopreservation of ovarian tissue for fertility preservation in
young female oncological patients. Future Oncol, 2012. 8(5): p. 595-608.
292. Eppig, J.J. and M.J. O'Brien, Development in vitro of mouse oocytes from primordial
follicles. Biol Reprod, 1996. 54(1): p. 197-207.
293. O'Brien, M.J., J.K. Pendola, and J.J. Eppig, A revised protocol for in vitro development of
mouse oocytes from primordial follicles dramatically improves their developmental
competence. Biol Reprod, 2003. 68(5): p. 1682-6.
294. Telfer, E.E. and M.B. Zelinski, Ovarian follicle culture: advances and challenges for
human and nonhuman primates. Fertil Steril, 2013. 99(6): p. 1523-33.
161
295. Cobo, A., et al., Is vitrification of oocytes useful for fertility preservation for age-related
fertility decline and in cancer patients? Fertil Steril, 2013. 99(6): p. 1485-95.
296. Bentov, Y., et al., The use of mitochondrial nutrients to improve the outcome of infertility
treatment in older patients. Fertil Steril, 2010. 93(1): p. 272-5.
297. Selesniemi, K., et al., Prevention of maternal aging-associated oocyte aneuploidy and
meiotic spindle defects in mice by dietary and genetic strategies. Proc Natl Acad Sci U S
A, 2011. 108(30): p. 12319-24.
298. Bender, T. and J.C. Martinou, Where killers meet--permeabilization of the outer
mitochondrial membrane during apoptosis. Cold Spring Harb Perspect Biol, 2013. 5(1):
p. a011106.
299. Noma, T., et al., Stage- and sex-dependent expressions of Usp9x, an X-linked mouse
ortholog of Drosophila Fat facets, during gonadal development and oogenesis in mice.
Mech Dev, 2002. 119 Suppl 1: p. S91-5.
300. Knauff, E.A., et al., Genome-wide association study in premature ovarian failure patients
suggests ADAMTS19 as a possible candidate gene. Hum Reprod, 2009. 24(9): p. 2372-8.
301. Davidson, J.S., I.M. Baumgarten, and E.H. Harley, Reversible inhibition of intercellular
junctional communication by glycyrrhetinic acid. Biochem Biophys Res Commun, 1986.
134(1): p. 29-36.
302. Pradelli, L.A., et al., Glycolysis inhibition sensitizes tumor cells to death receptors-
induced apoptosis by AMP kinase activation leading to Mcl-1 block in translation.
Oncogene, 2010. 29(11): p. 1641-52.
303. Ya, R. and S.M. Downs, Suppression of chemically induced and spontaneous mouse
oocyte activation by AMP-activated protein kinase. Biol Reprod, 2013. 88(3): p. 70.
304. Sullivan, J.E., et al., Inhibition of lipolysis and lipogenesis in isolated rat adipocytes with
AICAR, a cell-permeable activator of AMP-activated protein kinase. FEBS Lett, 1994.
353(1): p. 33-6.
305. Wu, X., et al., Involvement of AMP-activated protein kinase in glucose uptake stimulated
by the globular domain of adiponectin in primary rat adipocytes. Diabetes, 2003. 52(6):
p. 1355-63.
306. Dumont, F.J. and Q. Su, Mechanism of action of the immunosuppressant rapamycin. Life
Sci, 1996. 58(5): p. 373-95.
307. Thomas, R.L. and A.B. Gustafsson, Mitochondrial Autophagy. Circ J, 2013.
308. Twig, G., et al., Fission and selective fusion govern mitochondrial segregation and
elimination by autophagy. EMBO J, 2008. 27(2): p. 433-46.
162
309. Takatsuka, C., et al., 3-methyladenine inhibits autophagy in tobacco culture cells under
sucrose starvation conditions. Plant Cell Physiol, 2004. 45(3): p. 265-74.
310. Hoppins, S., et al., The soluble form of Bax regulates mitochondrial fusion via MFN2
homotypic complexes. Mol Cell, 2011. 41(2): p. 150-60.
311. Srinivasan, M., F. Kalousek, and N.P. Curthoys, In vitro characterization of the
mitochondrial processing and the potential function of the 68-kDa subunit of renal
glutaminase. J Biol Chem, 1995. 270(3): p. 1185-90.
312. Striebel, H.M., et al., Mutational analysis of both subunits from rat mitochondrial
processing peptidase. Arch Biochem Biophys, 1996. 335(1): p. 211-8.
313. Manabe, N., et al., Regulation mechanism of selective atresia in porcine follicles:
regulation of granulosa cell apoptosis during atresia. J Reprod Dev, 2004. 50(5): p. 493-
514.
314. Manabe, N., et al., Role of cell death ligand and receptor system on regulation of
follicular atresia in pig ovaries. Reprod Domest Anim, 2008. 43 Suppl 2: p. 268-72.
315. Dalton, C.M., G. Szabadkai, and J. Carroll, Dynamic measurement of ATP in single
oocytes: Impact of stage of maturation and cumulus cells on ATP levels and rates of
consumption. J Cell Physiol, 2013.
163
APPENDIX
Figure A1. Histomorphometric Analyses of BimKO and Protection Against Radiation-
Induced Primordial Follicle Death.
(i) Histomorphometric analyses of neonatal (PN4) ovaries of Bim-/-
(BimKO) (green) and Bim+/+
(blue) to
determine total number of alive and apoptotic (red) primordial (left). Additionally, total number of
surviving and apoptotic primordial follicle number was quantitated 48hrs after γ-irradiation (0.5Gy)
(right). Values represent average primordial follicle numbers ± SEM, both healthy or apoptotic (red) per
genotype. (ii) Image (Image magnification =400X) displays ovarian section of non-irradiated BimKO (top
left) and Bim+/+
(top right), in addition to BimKO (bottom left) and Bim+/+
(bottom right), 48hrs post
radiation. Presence of apoptotic primordial follicles oocytes marked by red arrowheads. (iii) Western
Blot (WB) displaying total ovarian lysates of BimKO and Bim+/+
stained with anti-BIM antibody with
ACTIN used as an internal control.
164
165
Figure A2. Assessing Impact of γ-Irradiation on MCL-1 and BIM Expression and MCL-1-
BIM Interaction.
(A) Neonatal mice were γ-irradiated (1Gy) and collected at select timepoints after radiation to determine
impact on MCL-1 and BIM expression. (i) Quantitation of band intensity of total MCL-1 using Quantity
One Software, at various timepoints 1hr (n=9), 2hrs (n=4), 4hrs (n=3), 8hrs (n=1) and 24hrs (n=2) after
irradiation. MCL-1 band intensity was normalized to ACTIN, used as an internal control. (ii) Western
Blots (WB) displaying impact of irradiation on levels of MCL-1 and 3 isoforms of pro-apoptotic BIM on
PN4 ovaries, 3hrs after irradiation. ACTIN was used as an internal control. Values represent fold change
of quantitated total MCL-1 ± SEM per condition, normalized to ACTIN at same time-point. (B) Co-IPs
were performed on irradiated neonatal ovaries and non-irradiated controls to assess impact of γ-irradiation
on MCL-1-BIM interaction. (i) Co-IP to assess interaction of MCL-1 and BIM. Total ovarian lysates
were immuno-precipitated with anti-Mcl-1, run on 12% SDS-PAGE gel and transferred to PVDF
membrane. Co-IP’s were immuno-blotted with antibodies against pro-apoptotic Bcl-2 members, BOK,
BAX and BIM. Anti-TOM20, a mitochondrial outer membrane import receptor, was utilized to determine
whether MCL-1 pulldown resulted in non-specific pulldown of total mitochondria outer membrane. (ii)
Reverse pulldown Co-IPs were performed on total ovarian lysates, immuno-precipitated with anti-Bim
immuno-blotted with anti-MCL-1 to confirm results from MCL-1 pulldown. (iii) In PN4 mice ovaries,
representative of a large primordial follicle population, either non-irradiated,1hr or 3hrs post γ-irradiation
(1Gy) ovarian lysates were used for co-IPs with MCL-1 pulldowns. These blots were immuno-blotted
with antibodies against pro-apoptotic Bcl-2 members BAX and BIM, to determine putative variations in
interaction post radiation.
