The Ovary [3 ed.] 9780128132098, 0128132094

Table of contents :
Front Cover
The Ovary
Copyright
Contents
Contributors
Part I: The Ovarian Follicular Apparatus: Operational Characteristics
Chapter 1: Follicle Selection in Mammalian Ovaries
Introduction
The Ovarian Reserve
Mammalian Follicles and Folliculogenesis
Selection of Ovarian Follicles
Selection of Primordial Follicles
Selection for Death
Selection to Activate
Dynamic Changes in the Selection for Death or Activation of Primordial Follicles Over the Fertile Life Span
Intra- and Extragonadal Hormonal Control of Follicle Selection
Gonadotropin-Independent Follicle Growth-Prenatal to Antral Development
Gonadotropin-Dependent Follicle Growth and Preovulatory Follicle Selection
Oocyte Regulation of Follicle Selection
Oocytes Participate in Follicle Selection via Key Paracrine Messengers
GDF9 and BMP15 Regulate Follicle Development, Selection, and Ovulation Rate
Oocyte Paracrine Signals Regulate GC Lineage Differentiation
Impact of Oocyte-Secreted Factor Signaling on Cumulus Cell and Oocyte Function
Oocyte-Somatic Cell Bidirectional Communication Maintains Oocyte Meiotic Arrest
Conclusions
References
Further Reading
Chapter 2: Regulation of Follicle Formation and Development by Ovarian Signaling Pathways
Introduction
Cellular Signals Regulating Germ Cell Nest Breakdown and Primordial Follicle Formation
Molecular Control of Neonatal Oocyte Survival
The Role of Steroid Hormones in Nest Breakdown
The Roles of Growth Factors and Somatic Cell-Cycle Progression During Germ Cell Nest Breakdown
Regulation of Germ Cell Nest Breakdown by Notch Signaling
Regulation of Primordial Follicle Activation
Intracellular Signaling Pathways That Regulate Primordial Follicle Activation
Regulation of Oocyte Activation by the PI3K/AKT Pathway
Contribution of mTOR Signaling to Primordial Follicle Activation
Oocyte-Specific Transcriptional Networks That Promote Primordial Follicle Activation
Granulosa Cell Differentiation During the Primordial to Primary Follicle Transition
Cellular Signaling During Preantral Follicular Development
TGF-β Family Signal Transduction in the Ovary
Control of Follicular Growth by Oocyte-Secreted GDF9 and BMP15
Regulation of Granulosa Cell Proliferation by Granulosa Cell-Derived TGF-βFamily Ligands
The Role of Notch Signaling in Preantral Follicle Development
Molecular Mechanisms of Theca Cell Recruitment and Specification
Regulation of Granulosa Cell Proliferation and Differentiation in Antral and Preovulatory Follicles
Crosstalk Between FSHR Signaling and Kinase Cascades in Granulosa Cell Proliferation and Differentiation
Control of Granulosa Cell Proliferation and Gene Expression by MAPK/ERK Activation Downstream of FSHR Signaling
Regulation of Granulosa Cell Differentiation and Proliferation by the PI3K/AKT Pathway
The Roles of Steroid Hormones in Follicular Maturation
Estrogen and Estrogen Receptor Signaling in Antral and Preovulatory Follicles
Physiological and Supraphysiological Effects of Androgens on Follicular Development
Conclusions
Supporting Grants
Disclosure Summary
References
Chapter 3: Human Folliculogenesis Revisited: The Menstrual Cycle Visualized by Ultrasonography
Introduction
Folliculogenesis
The Language of Folliculogenesis
Preantral and Early Antral Follicle Development
The Prepubertal and Pubertal Period
Recruitment of Antral Follicles
Folliculogenesis During the Interovulatory Interval
Dominant Follicle Selection
Follicle Divergence
Unified Theory of Selection
Follicle Dominance
Luteal Influences on Follicle Development and Selection
Preovulatory Follicle Development
Follicle Selection in Anovulatory Waves
Repeatability in Follicular Wave Patterns
Transition to Reproductive Senescence
Hormonal Influences on Folliculogenesis
Ovulation and Luteogenesis
Insights Regarding Animal Models for the Study of Folliculogenesis in Women
Future Directions and Clinical Implications
Summary
References
Chapter 4: Development of the Mammalian Ovary and Follicles
Introduction
Gonadal (or Genital) Ridge Formation
Ovigerous Cord Formation
Regionalization-Cortex and Medulla
Ovigerous Cord Breakdown and Follicle Formation
Establishment of Primordial Follicle Reserve
Follicle Activation
Formation of the Ovarian Surface Epithelium
Formation of the Ovarian Tunica Albuginea
Formation of the Theca Interna and Externa
Early Formation of the Vasculature
Conclusions
References
Chapter 5: Organization of Ovarian Steroidogenic Cells and Cholesterol Metabolism
Introduction
Organization of Steroidogenic Tissues and Cells
Acquisition, Storage, and Trafficking of Cholesterol
Plasma Membrane Cholesterol
Plasma Lipoproteins: The LDL and HDL Pathways
De Novo Cholesterol Synthesis
Relative Roles of Plasma Lipoproteins and De Novo Synthesis in Supplying Cholesterol for Steroidogenesis
Lipid Droplets
Intracellular Cholesterol Trafficking
Regulation of Cellular Cholesterol Balance
Overview of Steroidogenesis
Key Proteins in the Biosynthesis and Metabolism of Steroid Hormones
Steroidogenic Acute Regulatory Protein (STARD1): The Principal Regulator of Gonadal Steroidogenesis
The Cholesterol Side-Chain Cleavage Enzyme (P450scc Encoded by CYP11A1)
3β-Hydroxysteroid Dehydrogenase/Delta5-4 Isomerase
Regulation of Expression of the Steroidogenic Machinery
Conclusion
References
Further Reading
Chapter 6: Inhibin, Activin, and Follistatin in Ovarian Physiology
Biochemistry and Molecular Biology of Inhibin, Activin, and Follistatin
Structural Features, Signaling, and Regulation of Inhibin, Activin, and Follistatin Activity
Synthesis and Action of Inhibin, Activin, and Follistatin in the Ovary
Human Physiology and Clinical Implications
FSH Regulation for Controlled Follicle Development
Ovarian Aging, Perimenopause and Menopause
Primary Ovarian Insufficiency
Puberty
Pregnancy and Labor
In Vitro Fertilization and Ovarian Follicle Reserve
Ovarian Cancer
Polycystic Ovary Syndrome
Summary
References
Chapter 7: Cell-Cell Interactions in Ovarian Follicles: Role of TGF-β Superfamily Members
Introduction
TGF-β Superfamily Ligands
TGF-β Signaling Receptors
TGF-β Signal Transduction Pathway
Expression of TGF-β Superfamily Ligands in the Human Ovary
Expression of TGF-β Receptors in the Human Ovary
Primordial Germ Cell Development
Regulation of Ovarian Preantral Follicle Development
Intraovarian Cell-Cell Communication
Oocyte-Somatic Cell Interactions
Ovarian Steroidogenesis
GC Proliferation and Differentiation
Modulation of Extracellular Matrix Formation
Modulation of Cumulus-Oophorus Complex Formation and Expansion
Modulation of Ovulation
Modulation of Luteal Function
Roles of TGF-β Superfamily Members in Female Reproductive Pathology
Primary Ovarian Insufficiency
Polycystic Ovary Syndrome
Endometriosis
Perspectives for Therapeutic Development
Conclusion
Acknowledgment
References
Chapter 8: Mutations and Polymorphisms, and Their Functional Consequences, in Gonadotropin and Gonadotropin Receptor Genes
Introduction
Structural Features of Gonadotropins and Gonadotropin Receptors
Gonadotropins
Gonadotropin Subinit Genes and Proteins
Three-Dimensional Structure of hCG and FSH
Gonadotropin Receptors
Luteinizing Hormone/Chorionic Gonadotropin Receptor
LHCGR Gene, Messenger Ribonucleic Acid, and Protein
Activation and Signal Transduction of the LHCGR
Follicle-Stimulating Hormone Receptor
The FSHR Gene, Messenger Ribonucleic Acid, and Protein
Activation and Signal Transduction of the FSHR
Effects of Mutations and Polymorphisms in Gonadotropin and Gonadotropin Receptor Genes on Ovarian Function
Gonadotropin Subunits
Common α-Subunit (CGA)
LHB Subunit (LHB)
Mutations
Polymorphisms
hCG β-Subunit (CGB)
FSH β-Subunit (FSHB)
Mutations
Polymorphisms
Gonadotropin Receptors
Luteinizing Hormone/Chorionic Gonadotropin Receptor
Activating Mutations
Inactivating Mutations
Polymorphisms
FSHR
Activating Mutations
Inactivating Mutations
Polymorphisms
Conclusion
References
Chapter 9: Environmentally Induced Epigenetic Transgenerational Inheritance of Ovarian Disease
Introduction
The Epigenome
Transgenerational Ovarian Effects
Vinclozolin
Pesticides: Permethrin and DEET
Pesticide: DDT
Pesticide: Methoxychlor
Plastic Derived Compounds: BPA, DBP, and DEHP
Dioxin
Jet fuel: Hydrocarbon (JP8)
Conclusion
References
Part II: Oocyte Maturation and Ovulation
Chapter 10: Mammalian Oogenesis: The Fragile Foundation of the Next Generation
Introduction
Embryonic Development of the Ovarian Reserve
Cytoplasmic Maturation
Maternal Effect Genes
Cytoplasmic Reorganization
Nuclear Maturation
Epigenetic Regulation of the Genome
Conclusions
References
Chapter 11: Regulation of Mammalian Oocyte Maturation
Introduction
Nuclear Maturation
Key Cell Cycle Molecules Driving Nuclear Maturation
Meiotic Competence
Maintenance of Prophase Arrest in Oocytes From Antral Follicles
cAMP Produced in the Oocyte Promotes Prophase Arrest
cGMP From Granulosa Cells Regulates cAMP Levels Within the Oocyte
The Induction of Nuclear Maturation by LH
Epigenetic Maturation
Establishment of Epigenetic Modifications During Oogenesis
Large-Scale Chromatin Remodeling and Global Transcriptional Silencing in the Oocyte Genome
Conclusion
Acknowledgments
References
Chapter 12: Oocyte Meiotic Maturation
Introduction
Oocyte Nuclear Maturation
Mechanism of Oocyte Meiosis Arrest at Prophase I
Maintaining a High cAMP Level in the GV Oocyte
Maintaining a Low MPF Activity by cAMP in the GV Oocyte
Regulation of Cyclin B1 Levels
Balancing Protein Phosphatases
Oocyte Meiotic Resumption
Completion of the First Meiotic Division
Homologous Chromosome Separation
Interkinesis
Maintenance of MII Arrest
Oocyte Cytoplasmic Molecular Maturation
Translational Activation of Dormant mRNAs
Maternal mRNA Decay
BTG4 and CCR4-NOT RNA Deadenylase
Zinc Finger Protein 36 Like 2
Zygotic Arrest-1 (ZAR1) and -2 (ZAR2)
Oocyte Cytoplasmic Organelle Maturation
Mitochondrial Number and Distribution
Cortical Granule Migration
Redistribution of ER and Golgi Complex
Cytoskeleton Dynamics
Evaluation of Cytoplasmic Quality and Improvement of Cytoplasmic Maturation
Oocyte Epigenetic Maturation
Epigenetic Modifications in the GV Oocyte
Exchanges of Histone Variants During Oocyte Maturation
Histone H3 Lysine-4 Trimethylation
DNA Methylation and Demethylation in Oocytes
Polycomb-Repressive Complexes
Conclusion
Acknowledgments
References
Chapter 13: Gene Expression During Oogenesis and Oocyte Development
Introduction
Gene Expression During Oogenesis
Molecular Regulation of Oogenesis
Gene Expression During PGCs Migration and Proliferation
Gene Expression During Primordial Follicle Formation
Gene Expression During Follicle Activation and Development
Gene Expression Profile During Oocyte Maturation
The Regulation of Gene Expression During Oocyte Maturation
mRNA Stability in Oocytes
Translation Regulation in Oocytes
miRNA in Oocytes
Oocyte Proteome
Epigenetic Dynamics During PGC and Oocyte Development
Epigenetic Events During PGC Development
Epigenetic Events During Oocyte Growth and Maturation
Abnormal Expression of Genes in Reproductive Diseases
Conclusion
References
Chapter 14: Ovulation: The Coordination of Intrafollicular Networks to Ensure Oocyte Release
Introduction
Ovulatory Intracellular Signaling Cascades
Extracellular-Regulated Kinases Erk1/2
CCAAT Enhancer Binding Proteins CEBPα/CEBPβ
Progesterone Receptor
Cumulus Oocyte Complex Expansion
Extracellular Matrix Production
Role of Granulosa Cell EGF-Like Factors
Oocyte Control of Ovulatory Pathways
Proteolytic and Angiogenic Tissue Remodeling
Protease Actions
Angiogenesis
Vasoconstriction and Muscle Contraction
Immune Cell Infiltration and Inflammatory Mediators
Leukocyte Infiltration
Nitric Oxide and Reactive Oxygen Species
Prostaglandins
Cytokines
Clinical Aspects of Ovulation: Anovulation and Contraception
Diverse Etiology of Anovulation in Women
Contraceptives Targeting Ovulation
Summary and Conclusions
References
Part III: The Corpus Luteum
Chapter 15: Molecular Regulation of Progesterone Production in the Corpus Luteum
Introduction
Corpus Luteum
Cholesterol Ester Storage and Signals Regulating Cholesterol Availability
Storage of Cholesterol in Lipid Droplets
Hormone-Sensitive Lipase
AMP-Activated Protein Kinase
Inhibition of AMPK by LH
Activation of AMPK by PGF2α
Regulation of Genes Encoding Steroidogenic Pathway Proteins and Cholesterol Availability
STAR
Cytochrome P450 Side-Chain Cleavage Complex-FDX1, FDXR, and CYP11A1
FDX1
FDXR
CYP11A1
HSD3B1/2
LDLR and SCARB1
HMGCR
LIPE and NCEH1
Conclusion
References
Chapter 16: Corpus Luteum Formation
Introduction
Hypoxia and Corpus Luteum Formation
The Presence and Activation of HIF1A in the Corpus Luteum
HIF1 Activation in Hypoxia
Luteal HIF1A-Dependent Genes
Hypoxia-Independent Regulation of HIF1A
Role of microRNA-210 in Hypoxic Responses
Luteal Angiogenesis
Proangiogenic Factors: VEGFA, FGF2, and PGE2
Role of PGE2 in Luteal Angiogenesis
Matrix Remodeling
Conclusions
Glossary
References
Chapter 17: Luteolysis and the Corpus Luteum of Pregnancy
Introduction
Acquisition of Luteolytic Capacity
Initiation of Luteolysis
Specific Luteolytic Actions of PGF2A on the Diverse Cell Types of the CL
Actions of PGF2A on Steroidogenic Cells
Actions of PGF2A on Endothelial Cells
Specific Actions of Immune Cells and Cytokines in Luteolysis
Immune cells
Cytokines
Additional Molecular Mediators of Luteolysis
Cell Loss During Luteolysis
Luteal Rescue
Luteal Rescue in the Ruminant
Antiluteolytic Actions of IFNT on the Uterus
Direct Actions of IFNT on the CL That May Render It Less Sensitive to PGF2A
Luteal Rescue in Other Species
Primate
Rodents
Other Species
Other Changes in the CL During Early Pregnancy
Additional Potential Regulators of the CL in Early Pregnancy
Another Pivotal Maternal Recognition Period?
Conclusion
References
Part IV: Novel Experimental Models
Chapter 18: Transgenic Mouse Models in the Study of Ovarian Function
Introduction
Ovarian Differentiation and Germ Cell Development
Early Folliculogenesis: Primordial Follicle Activation to Follicle Formation
Antral and Preovulatory Follicle Development
Meiotic Arrest of Oocytes During Folliculogenesis and Resumption of Meiosis
Ovulation
Conclusion
Acknowledgments
References
Chapter 19: Genome-Wide Association Studies of Ovarian Function Disorders
Introduction
Polycystic Ovary Syndrome
The Two Large-Scale GWASs of PCOS in the Han Chinese Population
The GWASs of PCOS in Korean Populations
The GWASs of PCOS in European Populations
The Metaanalysis of PCOS-GWASs Across Ethnicities
Premature Ovarian Failure/Insufficiency
Genetic Risk Factors to POF/I
The First GWAS of POF/I in a Dutch Population
The Largest GWASs of POF/I in a Han Chinese Population
Other GWASs of POF/I Across Ethnicities
Cytogenomic (Comparative Genomic Hybridization, CGH Array) Studies of POF/I
Conclusion
Glossary
References
Chapter 20: Mitochondria Research in Human Reproduction
Introduction
Role of mtDNA Evolutionary Utility in Assisted Reproduction
Clinical Relevance Between mtDNA and Human Embryo Viability
Effect of Long-Acting GnRH Agonist Treatment on Mitochondrial Function in Endometriosis
Relationship Between Low Oxygen Tension and Mitochondrial Function in Embryo Development
Conclusion
References
Chapter 21: The Role of Mitochondria in Reproductive Function and Assisted Reproduction
The origins
Mitochondrial Segregation
Mitochondria in Oocytes
Mitochondrial Therapy
Reproductive Aging
Coenzyme Q10 Supplementation
Mitochondrial Transfer
Ooplasmic Transfer in Mature Human Oocytes
Use of Autologous Mitochondria From Ovarian Egg Precursor Cells
The Amount of Mitochondrial DNA in the Oocytes and Cumulus Cells
The Amount of mtDNA at the Cleavage Stage
The Amount of Mitochondrial DNA in the Blastocyst
Conclusion
References
Part V: Human Ovarian Pathophysiology: Select Aspects
Chapter 22: Ovarian Hyperstimulation Syndrome
Definition and Prevalence
Pathogenesis
Risk Factors
Before COS Begins
During COS
After Ovum Pickup
Clinical Manifestations
Diagnosis
Prevention
Before COS Begins
Selecting the Stimulation Protocol
Adjuvant Therapies
Aspirin
Metformin
During COS
Coasting
Ovulation Triggering With a GnRH Agonist
Lowering or Withholding hCG
Kisspeptin (Kp)
After Egg Retrieval
Oocyte/Embryo Cryopreservation
Dopamine Agonists
Albumin
Calcium
Letrozole
GnRH Analogs in the Luteal Phase
Combination of Treatments
Management
Mild OHSS
Moderate OHSS
Severe and Critical OHSS
Conclusions
References
Chapter 23: The Role of GnRH Agonist Triggering in GnRH Antagonist-Based Ovarian Stimulation Protocols
Introduction
The Mid-Cycle Ovulation Trigger in Normal Physiology
hCG-Based Triggering in Ovarian Stimulation Cycles
The Rational for GnRHa Triggering
The Benefit of Dual (LH and FSH) Triggering
GnRHa Triggering After Ovarian Stimulation With GnRH Antagonist Cotreatment
Number of Mature Oocytes Retrieved and Embryo Quality
GnRHa Triggering for Preventing OHSS
Pre-GnRH Antagonist Era
GnRH Antagonist Era
Luteolysis Post-GnRHa Triggering
LPS After Ovulation Triggering
Intense LPS
Modified Luteal Support With hCG
Modified Luteal Support With LH
Modified Luteal Support With GnRHa
Individualized Modified Luteal Support According to the Ovarian Response
GnRHa Trigger and "Freeze All"
Conclusions
Glossary
References
Chapter 24: The Ovarian Factor in Assisted Reproductive Technology
Definitions
Definition of Ovarian Factor
Definition of ART
Ovarian Reserve
Physiology of Ovarian Aging
Diagnosing Normal vs Abnormal OR
Why OR Determines Ovarian Stimulation
Definition of Treatment Success in ART
What Controls the Ovary?
The Hypothalamic Pituitary Axis
The Newly Discovered Adrenal-Ovarian Axis
The Immune System
How the Ovary Controls Treatment Success in IVF
Quantity and Quality of Oocytes
What Constitutes Oocyte Quality
Morphologically
Chromosomally
Affecting Ovarian Performance
Interventions Into the Gonadotropin Sensitive Stage of Folliculogenesis
Natural Cycle IVF
Mild Stimulation
Standard Stimulation Protocols
Niche Protocols
Interventions Into Earlier Stages of Folliculogenesis
Androgen Supplementation
HGH Supplementation
Timing of Oocyte Retrieval
In Vitro Management of Oocytes and Embryos
Conclusions and the Future
Conflict of Interest
References
Chapter 25: The Role of Antimullerian Hormone in Assisted Reproduction
Introduction
AMH in Ovarian Physiology
Inhibition of Primordial Follicle Recruitment
Inhibition of FSH Responsiveness
Factors Influencing AMH Levels
AMH Utility in ART
Age-Specific Normative AMH Values
Prognostication and Individualization of ART
Polycystic Ovarian Syndrome
Fertility Preservation
The AMH Assay: Potential Pitfalls and Solutions
Summary
Glossary
References
Chapter 26: Polycystic Ovary Syndrome
Introduction
Differential Diagnosis of Polycystic Ovarian Syndrome
Hypothyroidism
Hyperprolactinemia
Nonclassical Congenital Adrenal Hyperplasia
Androgen-Secreting Tumors
Cushing's Syndrome
Other Causes
Diagnosis and Clinical Stigmata of Polycystic Ovary Syndrome
Identification of Polycystic Ovaries
Hyperandrogenism
Hyperandrogenemia
Hirsutism
Chronic Anovulation
Insulin Resistance
Detection of Insulin Resistance in Women With Polycystic Ovary Syndrome
Oral Glucose Tolerance Testing and Hemoglobin A1C
Antimullerian Hormone and Polycystic Ovarian Syndrome
Clinical Sequelae of Polycystic Ovary Syndrome
Infertility and Chronic Anovulation
Gynecologic Cancer
Type 2 Diabetes Mellitus
Cardiovascular Disease
Treatment of PCOS
Generalized Treatment Strategies of Polycystic Ovary Syndrome
Lifestyle Modification
Ovarian Suppressive Therapies
Metformin
Thiazolidinediones
Treatment of Hirsutism
Treatment of Infertility
Lifestyle Modification
Clomiphene Citrate
Letrozole
Gonadotropins
Laparoscopic Ovarian Drilling
In Vitro Fertilization
Polycystic Ovary Syndrome: A Lifetime Disorder?
Genetic Etiology
Family Studies: Phenotypic Variation
Genome-Wide Association Studies and Next-Generation Sequencing in PCOS
Conclusion
References
Chapter 27: Genetic and Environmental Factors in the Etiology of Polycystic Ovary Syndrome
Introduction
Genetic Factors
Genome-Wide Association Studies
Chinese-Ancestry Studies
European-Ancestry Studies
Candidate Gene Studies
Steroid Production and Metabolism
Obesity
Beta-Cell Function
Ovarian Folliculogenesis
Epigenetic Modifications
Hypermethylation
Hypomethylation
Differential Methylation
X-Chromosome Inactivation
Micro-RNA
Environmental Factors
Fetal Programming Hypothesis
Animal-Based Studies
Human Observational Studies
Environmental Toxins
Triclocarban
Bisphenol A
Phthalates
Perfluoroalkyl Acids
Nicotine
Dietary-Related Weight Gain
Gut Microbiota
Lack of Exercise and Physical Activity
Physical Fitness
Reproductive Function
Cardiovascular Health
Psychological Well-Being
Socioeconomic Status
Advanced Glycation End Products
Conclusions
Acknowledgments
Conflicts of Interest
Financial Disclosure
References
Chapter 28: Germ Cell Failure and Ovarian Resistance: Human Genes and Disorders
Introduction
Embryology of Ovarian Development: Nature of Genes to Be Postulated
Ovarian Failure as a Result of Haploinsufficiency for the X Chromosome (X-Monosomy and X-Deletions)
Nomenclature
Monosomy X
X Short-Arm Deletions and Ovarian Failure
Candidate Genes for Xp Gonadal Determinants
X Long-Arm Deletions
Candidate Genes for Xq Gonadal Determinants
Ovarian Failure as a Result of Mendelian Causes
XX Gonadal Dysgenesis Without Somatic Anomalies
NOBOX (POF5)
FIGLA (POF6)
NR5A1 (POF7)
STAG3 (POF8)
HFM1 (POF9)
MCM8 (POF10)
CSB-PGBD3 (POF11)
SYCE1 (POF12)
MSH5 (POF13)
Perrault Syndrome (XX Gonadal Dysgenesis With Neurosensory Deafness)
Cerebellar Ataxia and XX Gonadal Dysgenesis
XX Gonadal Dysgenesis in Other Unique Malformation Syndromes
Pleiotropic Mendelian Disorders Showing Ovarian Failure
Germ-Cell Failure in Male (46,XY) and Female (46,XX) Sibs
Galactosemia
Carbohydrate-Deficient Glycoprotein (Phosphomannomutase Deficiency)
Deficiency of 17a-Hydroxylase/17,20-Desmolase Deficiency (CYP17)
Aromatase Mutations (CYP 19)
46,XX Agonadia
Fragile X Syndrome and Expansion of Triplet Nucleotide Repeats (CGG)
Myotonic Dystrophy and Expansion of Triplet Repeats (CTG)
Blepharophimosis-Ptosis-Epicanthus (FOXL2)
Autosomal-Dominant POF
Polygenic Factors in POF: Normal Continuations Variation as a Cause of Heritable EM
Ovarian Resistance
XX Gonadal Dysgenesis Due to FSHR Gene Mutation (C566T)
Inactivating Luteinizing Hormone Receptor Defect (46,XX)
References
Chapter 29: Environmental Contaminants and Ovarian Toxicity
Introduction
Biomonitoring Studies and Evidence of Ovarian Exposure
Epidemiological Evidence of Ovarian Dysfunction
Biological Plausibility of Effects and Potential Mechanisms
Summary and Conclusions
References
Chapter 30: Autologous Transplantation of Human Ovarian Tissue
Introduction
A Brief History of Ovarian Tissue Cryopreservation and Transplantation
Transplantation Techniques of Frozen-Thawed Ovarian Tissue
Orthotopic Ovarian Transplantation
Heterotopic Ovarian Transplantation
Current Success Rate of Ovarian Transplantation
Limitations With Ovarian Transplantation
Risk of Cancer Cell Reimplantation
Conclusion and Future Directions
Glossary
References
Chapter 31: Oncofertility: Preservation of Ovarian Function After a Cancer Diagnosis
Introduction
Minimizing the Impact of Cancer Therapy on the Ovary
Assessment of the Ovarian Reserve
Fertility Preservation Options in the Cancer Setting
Before Cancer Therapy
Embryo and Oocyte Cryopreservation
Ovarian Tissue Cryopreservation and Transplantation
After Cancer Therapy
Donor Oocytes and Embryos
Gestational Carrier or Adoption
Experimental Fertility Preservation Methods
Contraception and Menstrual Suppression for Oncology Patients
Conclusion
Glossary
Acknowledgments
References
Part VI: Human Ovarian Cancer
Chapter 32: Ovarian Cancers: Their Varied Origins and Pathologically Implicated Microenvironment
Introduction
Development of the Ovary and the Mullerian Ducts
Subtypes and Origins of Ovarian Cancer
The Ovarian Epithelial Cancers
Subtypes of Ovarian Epithelial Cancers
High-Grade Serous Carcinomas
Low-Grade Serous Carcinoma
Endometrioid Carcinoma
Clear Cell Carcinoma
Mucinous Carcinoma
Sex Cord-Stromal Tumors
Adult Granulosa Cell Tumors
Sertoli-Leydig Cell Tumors
Conclusions and Questions Arising
The Ovarian Cancer Microenvironment: A Preliminary Overview
Multifaceted Functions of the Ovarian Cancer Microenvironment in Disease Progression
Cancer-Associated Fibroblasts
Mesenchymal Stem Cells
Ovary-Associated Adipocytes
Angiogenesis and Neovasculature
Ovarian Cancer-Associated EVs
Concluding Remarks and Future Directions
Acknowledgments
References
Chapter 33: Phenotypic Plasticity and the Origins and Progression of Ovarian Cancer
Introduction
Phenotypic Plasticity in Ovarian Epithelial Cells
The Epithelial-to-Mesenchymal Transition
The Epithelial-to-Mesenchymal Transition and Stemness
Stemness in the Ovarian Surface Epithelium
Identification and Characterization of Stemness
Regulation of OSE Stemness
Stemness in the Fallopian Tube Epithelium
Phenotypic Plasticity in Ovulatory Wound Repair
Reepithelialization
Inflammation
Potential Consequences of Cell Plasticity
Heterogeneity of Ovarian Cancer
Consequences of the EMT on Ovarian Cancer Progression
Metastasis
Chemoresistance
Cancer Stem Cells
Immunosuppression
A Therapeutic Outlook: Targeting Plasticity
Does Plasticity Actually Play a Role in Tumor Progression?
Conclusion
References
Chapter 34: Novel Therapeutic Approaches and Targets for Ovarian Cancer
Introduction
Molecular Aberrations in Ovarian Cancer
Homologous Recombination and Poly-ADP-Ribose Polymerase Inhibitors
Olaparib
Rucaparib
Niraparib
Veliparib
Talazoparib
Targeting Angiogenesis: Vascular Endothelial Growth Factor and VEGF Receptors
Phosphatidylinositol-3-Kinase/AKT Pathway
Retrovirus-Associated DNA Sequences/v-raf 1 Murine Leukemia Viral Oncogene Homolog 1 Pathway/MAPK-ERK Kinase/Extracellular ...
EGFR Pathway Targets
Folate Receptor-Alpha
P53
Antimesothelin Antibodies
Notch Pathway
Aurora Kinase
SWItch/Sucrose Nonfermentable (SWI/SNF) Chromatin-Remodeling Complex
Histone Acetyltransferases and Histone Deacetylases
Cyclin-Dependent Kinases
SRC-Family Kinases
Platelet-Derived Growth Factor
Immunotherapy
Combination Therapy
Combination and Emerging Antiangiogenic Therapies
PI3K/AKT and MEK Inhibitor Combination Therapies
Side Effects for Targeted Therapies
Summary
Acknowledgments
Conflicts of Interest
References
Chapter 35: Molecular and Cellular Basis of Chemoresistance in Ovarian Cancer
Introduction
Mechanism of Cisplatin Action and Chemoresistance
The Tumor Microenvironment
Cellular Mechanisms in the Ovarian Cancer Cell Chemoresistance
Gelsolin and Chemoresistance
P53 and Chemoresistance
Inhibitor of Apoptosis Proteins and Chemoresistance
Hexokinase II and Chemoresistance
The PI3K/Akt Pathway
PI3-K/Akt Pathway in Ovarian Cancer Chemoresistance
The Interaction of Akt Signaling With XIAP, p53, HKII, and Gelsolin
Tumor Microenvironment and Chemorsistance
Effect of Chemotherapy on TILs and Implications for Immunotherapy
Cross-Talk Molecules in the Tumor Microenvironment
Tumor-Derived Extracellular Vesicles
Cancer-Derived Extracellular Vesicles and Chemoresistance
Plasma Gelsolin and Chemoresistance
Cytokines/Chemokines and Chemoresistance
Modern Treatment Strategies for Ovarian Chemoresistance
Immune Checkpoint Blockers
Cell-Based Immunotherapies
Targeting Epigenetic Changes
Natural Food Compounds
Nanotechnology/Nanomedicine
Personalized Cancer Therapy: The Future?
Conclusion
Glossary
References
Index
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THE OVARY THIRD EDITION

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THE OVARY THIRD EDITION Edited by

PETER C.K. LEUNG, PHD, FRSC, FCAHS, FKAST Professor of Obstetrics & Gynecology, Faculty of Medicine, University of British Columbia, Vancouver, BC, Canada

ELI Y. ADASHI, MD, MS, CPE, FACOG Professor of Medical Science The Warren Alpert Medical School, Brown University, Providence, RI, United States

Academic Press is an imprint of Elsevier 125 London Wall, London EC2Y 5AS, United Kingdom 525 B Street, Suite 1650, San Diego, CA 92101, United States 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, United Kingdom © 2019 Elsevier Inc. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. Library of Congress Cataloging-in-Publication Data A catalog record for this book is available from the Library of Congress British Library Cataloguing-in-Publication Data A catalogue record for this book is available from the British Library ISBN 978-0-12-813209-8 For information on all Academic Press publications visit our website at https://www.elsevier.com/books-and-journals

Publisher: John Fedor Acquisition Editor: Tari Broderick Editorial Project Manager: Kristi Anderson Production Project Manager: Sreejith Viswanathan Cover Designer: Miles Hitchens Typeset by SPi Global, India

Contents Contributors

Summary References

xi

I

4. Development of the Mammalian Ovary and Follicles

THE OVARIAN FOLLICULAR APPARATUS: OPERATIONAL CHARACTERISTICS

KATJA HUMMITZSCH, HELEN F. IRVING-RODGERS, JEFF SCHWARTZ, RAYMOND J. RODGERS

Introduction Gonadal (or Genital) Ridge Formation Ovigerous Cord Formation Regionalization—Cortex and Medulla Ovigerous Cord Breakdown and Follicle Formation Establishment of Primordial Follicle Reserve Follicle Activation Formation of the Ovarian Surface Epithelium Formation of the Ovarian Tunica Albuginea Formation of the Theca Interna and Externa Early Formation of the Vasculature Conclusions References

1. Follicle Selection in Mammalian Ovaries J.K. FINDLAY, K.R. DUNNING, R.B. GILCHRIST, K.J. HUTT, DARRYL L. RUSSELL, K.A. WALTERS

Introduction Selection of Primordial Follicles Intra- and Extragonadal Hormonal Control of Follicle Selection Oocyte Regulation of Follicle Selection Conclusions References Further Reading

3 5 8 11 17 17 21

JEROME F. STRAUSS, III

REXXI D. PRASASYA, KELLY E. MAYO

Introduction Organization of Steroidogenic Tissues and Cells Acquisition, Storage, and Trafficking of Cholesterol Plasma Membrane Cholesterol Plasma Lipoproteins: The LDL and HDL Pathways De Novo Cholesterol Synthesis Relative Roles of Plasma Lipoproteins and De Novo Synthesis in Supplying Cholesterol for Steroidogenesis Lipid Droplets Intracellular Cholesterol Trafficking Regulation of Cellular Cholesterol Balance Overview of Steroidogenesis Key Proteins in the Biosynthesis and Metabolism of Steroid Hormones Regulation of Expression of the Steroidogenic Machinery Conclusion References Further Reading

23 24 29 34 38 42 42 42 42

3. Human Folliculogenesis Revisited: The Menstrual Cycle Visualized by Ultrasonography ROGER A. PIERSON

Introduction Folliculogenesis Hormonal Influences on Folliculogenesis Ovulation and Luteogenesis Insights Regarding Animal Models for the Study of Folliculogenesis in Women Future Directions and Clinical Implications

71 72 75 76 76 76 77 77 78 78 78 79 79

5. Organization of Ovarian Steroidogenic Cells and Cholesterol Metabolism

2. Regulation of Follicle Formation and Development by Ovarian Signaling Pathways Introduction Cellular Signals Regulating Germ Cell Nest Breakdown and Primordial Follicle Formation Regulation of Primordial Follicle Activation Cellular Signaling During Preantral Follicular Development Regulation of Granulosa Cell Proliferation and Differentiation in Antral and Preovulatory Follicles Conclusions Supporting Grants Disclosure Summary References

63 64

51 51 60 61

6. Inhibin, Activin, and Follistatin in Ovarian Physiology

62 63

Biochemistry and Molecular Biology of Inhibin, Activin, and Follistatin

83 83 83 84 84 85 85 86 87 87 88 88 92 92 93 94

CORRINE WELT, ALAN SCHNEYER

v

95

vi Structural Features, Signaling, and Regulation of Inhibin, Activin, and Follistatin Activity Synthesis and Action of Inhibin, Activin, and Follistatin in the Ovary Human Physiology and Clinical Implications Summary References

CONTENTS

96 97 99 101 102

7. Cell-Cell Interactions in Ovarian Follicles: Role of TGF-β Superfamily Members HSUN-MING CHANG, YI-MIN ZHU, PETER C.K. LEUNG

Introduction TGF-β Superfamily Ligands TGF-β Signaling Receptors TGF-β Signal Transduction Pathway Expression of TGF-β Superfamily Ligands in the Human Ovary Expression of TGF-β Receptors in the Human Ovary Primordial Germ Cell Development Regulation of Ovarian Preantral Follicle Development Intraovarian Cell-Cell Communication Oocyte-Somatic Cell Interactions Ovarian Steroidogenesis GC Proliferation and Differentiation Modulation of Extracellular Matrix Formation Modulation of Cumulus-Oophorus Complex Formation and Expansion Modulation of Ovulation Modulation of Luteal Function Roles of TGF-β Superfamily Members in Female Reproductive Pathology Primary Ovarian Insufficiency Polycystic Ovary Syndrome Endometriosis Perspectives for Therapeutic Development Conclusion Acknowledgment References

107 107 108 108 109 111 112 112 113 113 115 116 116 116 117 117 118 118 119 119 119 120 120 120

8. Mutations and Polymorphisms, and Their Functional Consequences, in Gonadotropin and Gonadotropin Receptor Genes € ILPO HUHTANIEMI, ADOLFO RIVERO-MULLER

Introduction 127 Structural Features of Gonadotropins and Gonadotropin Receptors 127 Effects of Mutations and Polymorphisms in Gonadotropin and Gonadotropin Receptor Genes on Ovarian Function 132 Conclusion 144 References 145

9. Environmentally Induced Epigenetic Transgenerational Inheritance of Ovarian Disease H.J. KIMBEL, E.E. NILSSON, M.K. SKINNER

Introduction The Epigenome

149 151

Transgenerational Ovarian Effects Conclusion References

151 153 153

II OOCYTE MATURATION AND OVULATION 10. Mammalian Oogenesis: The Fragile Foundation of the Next Generation JOHN J. BROMFIELD, RACHEL L. PIERSANTI

Introduction Embryonic Development of the Ovarian Reserve Cytoplasmic Maturation Nuclear Maturation Epigenetic Regulation of the Genome Conclusions References

157 157 157 159 161 162 162

11. Regulation of Mammalian Oocyte Maturation MARIA M. VIVEIROS, RABINDRANATH DE LA FUENTE

Introduction Nuclear Maturation Epigenetic Maturation Conclusion Acknowledgments References

165 165 170 176 176 176

12. Oocyte Meiotic Maturation HENG-YU FAN, QING-YUAN SUN

Introduction Oocyte Nuclear Maturation Oocyte Cytoplasmic Molecular Maturation Oocyte Cytoplasmic Organelle Maturation Oocyte Epigenetic Maturation Conclusion Acknowledgments References

181 181 189 192 196 200 200 200

13. Gene Expression During Oogenesis and Oocyte Development MO LI, JIE YAN, XU ZHI, YUN WANG, JING HANG, JIE QIAO

Introduction Gene Expression During Oogenesis Gene Expression Profile During Oocyte Maturation The Regulation of Gene Expression During Oocyte Maturation Epigenetic Dynamics During PGC and Oocyte Development Abnormal Expression of Genes in Reproductive Diseases Conclusion References

205 205 208 209 211 213 214 214

vii

CONTENTS

IV

14. Ovulation: The Coordination of Intrafollicular Networks to Ensure Oocyte Release

NOVEL EXPERIMENTAL MODELS

DARRYL L. RUSSELL, REBECCA L. ROBKER

Introduction Ovulatory Intracellular Signaling Cascades Cumulus Oocyte Complex Expansion Proteolytic and Angiogenic Tissue Remodeling Vasoconstriction and Muscle Contraction Immune Cell Infiltration and Inflammatory Mediators Clinical Aspects of Ovulation: Anovulation and Contraception Summary and Conclusions References

217 218 221 223 224 225 226 228 229

III THE CORPUS LUTEUM 15. Molecular Regulation of Progesterone Production in the Corpus Luteum

16. Corpus Luteum Formation

237 238 239 244 248 248

255

KETAN SHRESTHA, DANIELA RODLER, FRED SINOWATZ, RINA MEIDAN

Introduction Conclusions References

AMANDA RODRIGUEZ, ROBERT T. RYDZE, SHAWN M. BRILEY, STEPHANIE A. PANGAS

Introduction Ovarian Differentiation and Germ Cell Development Early Folliculogenesis: Primordial Follicle Activation to Follicle Formation Antral and Preovulatory Follicle Development Meiotic Arrest of Oocytes During Folliculogenesis and Resumption of Meiosis Ovulation Conclusion Acknowledgments References

300 303 305 306 307 307 307

255 262 263

CHI-KWAN LEUNG, HAN ZHAO, YUE LV, GANG LU, ZI-JIANG CHEN

Introduction Polycystic Ovary Syndrome The Two Large-Scale GWASs of PCOS in the Han Chinese Population The GWASs of PCOS in Korean Populations The GWASs of PCOS in European Populations The Metaanalysis of PCOS-GWASs Across Ethnicities Premature Ovarian Failure/Insufficiency Genetic Risk Factors to POF/I The First GWAS of POF/I in a Dutch Population The Largest GWASs of POF/I in a Han Chinese Population Other GWASs of POF/I Across Ethnicities Cytogenomic (Comparative Genomic Hybridization, CGH Array) Studies of POF/I Conclusion References

17. Luteolysis and the Corpus Luteum of Pregnancy

20. Mitochondria Research in Human Reproduction

CAMILLA K. HUGHES, JOY L. PATE

YI-XUAN LEE, PEI-HSUAN LIN, ENDAH RAHMAWATI, YUN-YI MA, CINDY CHAN, CHII-RUEY TZENG

Introduction Acquisition of Luteolytic Capacity Initiation of Luteolysis Specific Luteolytic Actions of PGF2A on the Diverse Cell Types of the CL Specific Actions of Immune Cells and Cytokines in Luteolysis Additional Molecular Mediators of Luteolysis Cell Loss During Luteolysis Luteal Rescue Conclusion References

296 296

19. Genome-Wide Association Studies of Ovarian Function Disorders

JOHN S. DAVIS, HOLLY A. LAVOIE

Introduction Corpus Luteum Cholesterol Ester Storage and Signals Regulating Cholesterol Availability Regulation of Genes Encoding Steroidogenic Pathway Proteins and Cholesterol Availability Conclusion References

18. Transgenic Mouse Models in the Study of Ovarian Function

270 270 271 272 275 278 279 279 285 286

Introduction Role of mtDNA Evolutionary Utility in Assisted Reproduction Clinical Relevance Between mtDNA and Human Embryo Viability Effect of Long-Acting GnRH Agonist Treatment on Mitochondrial Function in Endometriosis Relationship Between Low Oxygen Tension and Mitochondrial Function in Embryo Development Conclusion References

311 311 311 317 317 318 318 319 319 319 319 321 322 323

327 327 329 330 331 333 333

viii

CONTENTS

21. The Role of Mitochondria in Reproductive Function and Assisted Reproduction JIGAL HAAS, RAWAD BASSIL, ROBERT F. CASPER

The origins Mitochondrial Segregation Mitochondria in Oocytes Mitochondrial Therapy Reproductive Aging Coenzyme Q10 Supplementation Mitochondrial Transfer Conclusion References

337 337 338 338 339 339 339 341 341

V HUMAN OVARIAN PATHOPHYSIOLOGY: SELECT ASPECTS

345 345 348 348 350 351 354 357 357

23. The Role of GnRH Agonist Triggering in GnRH Antagonist-Based Ovarian Stimulation Protocols SHAHAR KOL, OFER FAINARU

363 363 364 365 365 366 367 367 368 375 375

24. The Ovarian Factor in Assisted Reproductive Technology NORBERT GLEICHER, VITALY A. KUSHNIR, DAVID H. BARAD

Definitions Ovarian Reserve Definition of Treatment Success in ART What Controls the Ovary?

25. The Role of Antimullerian Hormone in Assisted Reproduction RESHEF TAL, DAVID B. SEIFER

Introduction AMH in Ovarian Physiology Factors Influencing AMH Levels AMH Utility in ART The AMH Assay: Potential Pitfalls and Solutions Summary References

403 403 405 406 409 409 410

IAN N. WALDMAN, RICHARD S. LEGRO

NURIA PELLICER, DANIELA GALLIANO, ANTONIO PELLICER

Introduction The Mid-Cycle Ovulation Trigger in Normal Physiology hCG-Based Triggering in Ovarian Stimulation Cycles The Rational for GnRHa Triggering The Benefit of Dual (LH and FSH) Triggering GnRHa Triggering After Ovarian Stimulation With GnRH Antagonist Cotreatment GnRHa Triggering for Preventing OHSS Luteolysis Post-GnRHa Triggering LPS After Ovulation Triggering Conclusions References

388 391 397 397 398

26. Polycystic Ovary Syndrome

22. Ovarian Hyperstimulation Syndrome Definition and Prevalence Pathogenesis Risk Factors Clinical Manifestations Diagnosis Prevention Management Conclusions References

How the Ovary Controls Treatment Success in IVF Affecting Ovarian Performance In Vitro Management of Oocytes and Embryos Conclusions and the Future References

379 381 383 385

Introduction Differential Diagnosis of Polycystic Ovarian Syndrome Hypothyroidism Hyperprolactinemia Nonclassical Congenital Adrenal Hyperplasia Androgen-Secreting Tumors Cushing’s Syndrome Other Causes Diagnosis and Clinical Stigmata of Polycystic Ovary Syndrome Identification of Polycystic Ovaries Hyperandrogenism Hyperandrogenemia Hirsutism Chronic Anovulation Insulin Resistance Detection of Insulin Resistance in Women With Polycystic Ovary Syndrome Oral Glucose Tolerance Testing and Hemoglobin A1C Antimullerian Hormone and Polycystic Ovarian Syndrome Clinical Sequelae of Polycystic Ovary Syndrome Generalized Treatment Strategies of Polycystic Ovary Syndrome Treatment of Infertility Polycystic Ovary Syndrome: A Lifetime Disorder? Genetic Etiology Family Studies: Phenotypic Variation Genome-Wide Association Studies and Next-Generation Sequencing in PCOS Conclusion References

415 415 415 415 416 416 416 417 417 417 418 418 418 419 419 420 420 421 421 423 425 427 428 428 428 429 429

27. Genetic and Environmental Factors in the Etiology of Polycystic Ovary Syndrome T.M. BARBER, S. FRANKS

Introduction Genetic Factors

437 438

ix

CONTENTS

Epigenetic Modifications Environmental Factors Fetal Programming Hypothesis Environmental Toxins Dietary-Related Weight Gain Gut Microbiota Lack of Exercise and Physical Activity Socioeconomic Status Advanced Glycation End Products Conclusions Acknowledgments References

441 443 443 445 447 449 450 451 452 454 455 455

461 461 462 465 480 480 481

29. Environmental Contaminants and Ovarian Toxicity 485 486 486 487 488 488

30. Autologous Transplantation of Human Ovarian Tissue ENES TAYLAN, KUTLUK OKTAY

493 493 494 497 497 498 498 499

31. Oncofertility: Preservation of Ovarian Function After a Cancer Diagnosis AZZA M. AKASHA, TERESA K. WOODRUFF

Introduction Minimizing the Impact of Cancer Therapy on the Ovary Assessment of the Ovarian Reserve

VI HUMAN OVARIAN CANCER

Introduction Development of the Ovary and the Mullerian Ducts Subtypes and Origins of Ovarian Cancer Conclusions and Questions Arising The Ovarian Cancer Microenvironment: A Preliminary Overview Multifaceted Functions of the Ovarian Cancer Microenvironment in Disease Progression Concluding Remarks and Future Directions Acknowledgments References

511 512 514 518 519 520 524 525 525

33. Phenotypic Plasticity and the Origins and Progression of Ovarian Cancer

W.G. FOSTER, A.M. GANNON, H.C. FURLONG

Introduction A Brief History of Ovarian Tissue Cryopreservation and Transplantation Transplantation Techniques of Frozen-Thawed Ovarian Tissue Current Success Rate of Ovarian Transplantation Limitations With Ovarian Transplantation Risk of Cancer Cell Reimplantation Conclusion and Future Directions References

504 506 506 506

YU SUN, NELLY AUERSPERG

JOE LEIGH SIMPSON, YINGYING QIN, ZI-JIANG CHEN

Introduction Biomonitoring Studies and Evidence of Ovarian Exposure Epidemiological Evidence of Ovarian Dysfunction Biological Plausibility of Effects and Potential Mechanisms Summary and Conclusions References

503

32. Ovarian Cancers: Their Varied Origins and Pathologically Implicated Microenvironment

28. Germ Cell Failure and Ovarian Resistance: Human Genes and Disorders Introduction Embryology of Ovarian Development: Nature of Genes to Be Postulated Ovarian Failure as a Result of Haploinsufficiency for the X Chromosome (X-Monosomy and X-Deletions) Ovarian Failure as a Result of Mendelian Causes Polygenic Factors in POF: Normal Continuations Variation as a Cause of Heritable EM Ovarian Resistance References

Fertility Preservation Options in the Cancer Setting Contraception and Menstrual Suppression for Oncology Patients Conclusion Acknowledgments References

501 501 502

LAUREN E. CARTER, DAVID P. COOK, BARBARA C. VANDERHYDEN

Introduction Phenotypic Plasticity in Ovarian Epithelial Cells The Epithelial-to-Mesenchymal Transition The Epithelial-to-Mesenchymal Transition and Stemness Stemness in the Ovarian Surface Epithelium Stemness in the Fallopian Tube Epithelium Phenotypic Plasticity in Ovulatory Wound Repair Heterogeneity of Ovarian Cancer Consequences of the EMT on Ovarian Cancer Progression A Therapeutic Outlook: Targeting Plasticity Conclusion References

529 529 530 531 532 532 534 536 537 540 540 541

34. Novel Therapeutic Approaches and Targets for Ovarian Cancer REBECCA A. PREVIS, GORDON B. MILLS, SHANNON N. WESTIN

Introduction 547 Molecular Aberrations in Ovarian Cancer 547 Homologous Recombination and Poly-ADP-Ribose Polymerase Inhibitors 549 Targeting Angiogenesis: Vascular Endothelial Growth Factor and VEGF Receptors 553 Phosphatidylinositol-3-Kinase/AKT Pathway 555 Retrovirus-Associated DNA Sequences/v-raf 1 Murine Leukemia Viral Oncogene Homolog 1 Pathway/MAPK-ERK Kinase/Extracellular Signal-Regulated Kinase 556

x EGFR Pathway Targets Folate Receptor-Alpha P53 Antimesothelin Antibodies Notch Pathway Aurora Kinase SWItch/Sucrose Nonfermentable (SWI/SNF) Chromatin-Remodeling Complex Histone Acetyltransferases and Histone Deacetylases Cyclin-Dependent Kinases SRC-Family Kinases Platelet-Derived Growth Factor Immunotherapy Combination Therapy Side Effects for Targeted Therapies Summary Acknowledgment References

CONTENTS

557 558 558 559 559 559 559 560 560 560 560 561 562 565 566 566 566

35. Molecular and Cellular Basis of Chemoresistance in Ovarian Cancer MESHACH ASARE-WEREHENE, DAR-BIN SHIEH, YONG SANG SONG, BENJAMIN K. TSANG

Introduction Mechanism of Cisplatin Action and Chemoresistance The Tumor Microenvironment Tumor Microenvironment and Chemorsistance Effect of Chemotherapy on TILs and Implications for Immunotherapy Cross-Talk Molecules in the Tumor Microenvironment Modern Treatment Strategies for Ovarian Chemoresistance Natural Food Compounds Nanotechnology/Nanomedicine Personalized Cancer Therapy: The Future? Conclusion References

Index

575 575 576 582 584 584 586 587 588 588 588 589

595

Contributors

Eli Y. Adashi Department of Medical Science, The Warren Alpert Medical School, Brown University, Providence, RI, United States

Cindy Chan Center for Reproductive Medicine, Department of Obstetrics and Gynecology, Taipei Medical University Hospital, Taipei, Taiwan

Azza M. Akasha Northwestern University, Department of Obstetrics and Gynecology, Feinberg School of Medicine, Chicago, IL, United States

Hsun-Ming Chang Department of Obstetrics and Gynaecology, BC Children’s Hospital Research Institute, University of British Columbia, Vancouver, BC, Canada

Meshach Asare-Werehene Departments of Obstetrics and Gynaecology and Cellular and Molecular Medicine, Interdisciplinary School of Health Sciences, University of Ottawa; Chronic Disease Program, Ottawa Hospital Research Institute, Ottawa, ON, Canada; State Key Laboratory of Quality Research in Chinese Medicine, Macau Institute for Applied Research in Medicine and Health, Macau University of Science and Technology, Taipa, Macao; Institute of Basic Medical Science, Institute of Oral Medicine and Department of Stomatology, National Cheng Kung University Hospital, Advanced Optoelectronic Technology Center & Center for Micro/Nano Science & Technology, College of Medicine, National Cheng Kung University, Tainan, Taiwan; Interdisciplinary Program in Cancer Biology, Cancer Research Institute, and the Department of Obstetrics and Gynecology, Seoul National University College of Medicine, Seoul, Republic of Korea

Zi-Jiang Chen National Research Center for Assisted Reproductive Technology and Reproductive Genetics; Center for Reproductive Medicine, Shandong University; The Key Laboratory for Reproductive Endocrinology, Shandong University, Ministry of Education, Jinan; Center for Reproductive Medicine, Ren Ji Hospital, School of Medicine, Shanghai Jiao Tong University; Shanghai Key Laboratory for Assisted Reproduction and Reproductive Genetics, Shanghai, China David P. Cook Department of Cellular and Molecular Medicine, University of Ottawa; Cancer Therapeutics Program, Ottawa Hospital Research Institute, Ottawa, ON, Canada John S. Davis Department of Obstetrics and Gynecology; Department of Biochemistry and Molecular Biology, University of Nebraska Medical Center; Veterans Affairs Nebraska-Western Iowa Health Care System, Omaha, NE, United States

Nelly Auersperg Department of Obstetrics and Gynecology, University of British Columbia, Vancouver, BC, Canada David H. Barad Center for Human Reproduction; Foundation for Reproductive Medicine, New York, NY, United States

Rabindranath De La Fuente Department of Physiology and Pharmacology, College of Veterinary Medicine; Regenerative Biosciences Center (RBC), University of Georgia, Athens, GA, United States

T.M. Barber Division of Biomedical Sciences (T.M.B.), Warwick Medical School, University of Warwick, Clinical Sciences Research Laboratories, University Hospitals Coventry and Warwickshire, Coventry, United Kingdom

K.R. Dunning Australian Research Council Centre of Excellence for Nanoscale BioPhotonics, Robinson Research Institute, University of Adelaide, Adelaide, SA, Australia

Rawad Bassil Division of Reproductive Sciences, University of Toronto, Lunenfeld-Tanenbaum Research Institute, Mount Sinai Hospital, Toronto, ON, Canada

Ofer Fainaru IVF Unit, Rambam Health Care Campus, Haifa, Israel

Shawn M. Briley Department of Pathology & Immunology; Graduate Program in Biochemistry & Molecular Biology, Baylor College of Medicine, Houston, TX, United States

Heng-Yu Fan Life Sciences Institute, Zhejiang University, Hangzhou, China J.K. Findlay Hudson Institute of Medical Research & Monash University, Clayton, VIC, Australia

John J. Bromfield Department of Animal Sciences, University of Florida, Gainesville, FL, United States

W.G. Foster Department of Obstetrics & Gynecology, McMaster University, Hamilton, ON, Canada

Lauren E. Carter Department of Cellular and Molecular Medicine, University of Ottawa; Cancer Therapeutics Program, Ottawa Hospital Research Institute, Ottawa, ON, Canada

S. Franks Institute of Reproductive and Developmental Biology (S.F.), Imperial College (Hammersmith Campus) London, United Kingdom H.C. Furlong Trinity College Dublin, The University of Dublin, Dublin, Ireland

Robert F. Casper Department of Obstetrics and Gynecology, University of Toronto; Lunenfeld-Tanenbaum Research Institute; TRIO Fertility; Insception-Lifebank Cord Blood Bank, Toronto, ON, Canada

Daniela Galliano Rome, Italy

xi

Instituto Valenciano de Infertilidad (IVI),

xii

CONTRIBUTORS

A.M. Gannon Food Directorate, Health Protection Branch, Health Canada, Ottawa, ON, Canada R.B. Gilchrist School of Women’s & Children’s Health, University of New South Wales, Randwick, NSW, Australia Norbert Gleicher Center for Human Reproduction; Foundation for Reproductive Medicine; Stem Cell Biology and Molecular Embryology Laboratory, The Rockefeller University, New York, NY, United States; Department of Obstetrics and Gynecology, University of Vienna School of Medicine, Vienna, Austria Jigal Haas Tel Hashomer Hospital, Tel Aviv University, Tel Aviv, Israel Jing Hang Center for Reproductive Medicine, Department of Obstetrics and Gynecology, Peking University Third Hospital, Beijing, China Camilla K. Hughes Center for Reproductive Biology and Health, Department of Animal Science, The Pennsylvania State University, Pennsylvania, PA, United States Ilpo Huhtaniemi Institute of Reproductive and Developmental Biology, Department of Surgery & Cancer, Imperial College London, London, United Kingdom; Department of Physiology, Institute of Biomedicine, University of Turku, Turku, Finland Katja Hummitzsch Discipline of Obstetrics and Gynaecology, School of Medicine, Robinson Research Institute, University of Adelaide, Adelaide, SA, Australia K.J. Hutt Biomedicine Discovery Institute, Department of Anatomy & Developmental Biology, Monash University, Clayton, VIC, Australia Helen F. Irving-Rodgers Discipline of Obstetrics and Gynaecology, School of Medicine, Robinson Research Institute, University of Adelaide, Adelaide, SA; School of Medical Science, Griffith University, Gold Coast, QLD, Australia H.J. Kimbel Center for Reproductive Biology, School of Biological Sciences, Washington State University, Pullman, WA, United States Shahar Kol

IVF Unit, Elisha Hospital, Haifa, Israel

Vitaly A. Kushnir Center for Human Reproduction, New York, NY; Department of Obstetrics and Gynecology, Wake Forest University, Winston Salem, NC, United States Holly A. LaVoie Department of Cell Biology & Anatomy, University of South Carolina School of Medicine, Columbia, SC, United States Yi-Xuan Lee Center for Reproductive Medicine, Department of Obstetrics and Gynecology, Taipei Medical University Hospital, Taipei, Taiwan Richard S. Legro Department of Obstetrics and Gynecology, Penn State College of Medicine, Hershey, PA, United States Peter C.K. Leung Department of Obstetrics & Gynecology, University of British Columbia, Vancouver, BC, Canada Chi-Kwan Leung National Research Center for Assisted Reproductive Technology and Reproductive Genetics; Center for Reproductive Medicine, Shandong University, Jinan; SDIVF R&D Centre, Hong Kong Science and Technology

Parks; CUHK-SDU Joint Laboratory on Reproductive Genetics, School of Biomedical Sciences, The Chinese University of Hong Kong, Hong Kong, China Mo Li Center for Reproductive Medicine, Department of Obstetrics and Gynecology, Peking University Third Hospital, Beijing, China Pei-Hsuan Lin Department of Obstetrics and Gynecology, Yuan’s General Hospital, Kaohsiung, Taiwan Gang Lu National Research Center for Assisted Reproductive Technology and Reproductive Genetics; Center for Reproductive Medicine, Shandong University, Jinan; CUHK-SDU Joint Laboratory on Reproductive Genetics, School of Biomedical Sciences, The Chinese University of Hong Kong, Hong Kong, China Yue Lv National Research Center for Assisted Reproductive Technology and Reproductive Genetics; Center for Reproductive Medicine, Shandong University; The Key Laboratory for Reproductive Endocrinology, Shandong University, Ministry of Education, Jinan; CUHK-SDU Joint Laboratory on Reproductive Genetics, School of Biomedical Sciences, The Chinese University of Hong Kong, Hong Kong, China Yun-Yi Ma Center for Reproductive Medicine, Department of Obstetrics and Gynecology, Taipei Medical University Hospital, Taipei, Taiwan Kelly E. Mayo Department of Molecular Biosciences and Center for Reproductive Science, Northwestern University, Evanston, IL, United States Rina Meidan Department of Animal Sciences, The Robert H. Smith Faculty of Agriculture, Food, and Environment, The Hebrew University of Jerusalem, Rehovot, Israel Gordon B. Mills Department of Systems Biology, University of Texas MD Anderson Cancer Center, Houston, TX, United States E.E. Nilsson Center for Reproductive Biology, School of Biological Sciences, Washington State University, Pullman, WA, United States Kutluk Oktay Laboratory of Molecular Reproduction and Fertility Preservation, Department of Obstetrics, Gynecology and Reproductive Sciences, Yale University School of Medicine, New Haven, CT, United States Stephanie A. Pangas Department of Pathology & Immunology; Graduate Program in Molecular & Cellular Biology; Clinical Scientist Training Program; Graduate Program in Biochemistry & Molecular Biology, Baylor College of Medicine, Houston, TX, United States Joy L. Pate Center for Reproductive Biology and Health, Department of Animal Science, The Pennsylvania State University, Pennsylvania, PA, United States Nuria Pellicer Department of Obstetrics and Gynecology, Hospital Universitario y Polit
ecnico, La Fe, Valencia, Spain Antonio Pellicer Instituto Valenciano de Infertilidad (IVI), Rome, Italy; Instituto de Investigación Sanitaria La Fe, Valencia, Spain Rachel L. Piersanti Department of Animal Sciences, University of Florida, Gainesville, FL, United States

CONTRIBUTORS

Roger A. Pierson Department of Obstetrics and Gynecology, College of Medicine, University of Saskatchewan, Saskatoon, Canada Rexxi D. Prasasya Department of Molecular Biosciences and Center for Reproductive Science, Northwestern University, Evanston, IL, United States Rebecca A. Previs Department of Obstetrics & Gynecology, Division of Gynecologic Oncology, Duke Cancer Institute, Duke University Medical Center, Durham, NC, United States Jie Qiao Center for Reproductive Medicine, Department of Obstetrics and Gynecology, Peking University Third Hospital, Beijing, China Yingying Qin Center for Reproductive Medicine, Shandong University, Ji’nan, China Endah Rahmawati Graduate Institute of Clinical Medicine, Taipei Medical University, Taipei, Taiwan; Department of Obstetrics and Gynecology, Faculty of Medicine, Universitas Gadjah Mada, Yogyakarta, Indonesia Adolfo Rivero-M€ uller Department of Physiology, Institute of Biomedicine, University of Turku, Turku, Finland; Department of Biochemistry and Molecular Biology, Medical University of Lublin, Lublin, Poland Rebecca L. Robker Robinson Research Institute, School of Medicine, University of Adelaide, Adelaide, SA, Australia Raymond J. Rodgers Discipline of Obstetrics and Gynaecology, School of Medicine, Robinson Research Institute, University of Adelaide, Adelaide, SA, Australia Daniela Rodler Department of Veterinary Sciences, LudwigMaximilian University Munich, Munich, Germany Amanda Rodriguez Department of Pathology & Immunology; Graduate Program in Molecular & Cellular Biology, Baylor College of Medicine, Houston, TX, United States Darryl L. Russell Robinson Research Institute, University of Adelaide, Adelaide, SA, Australia; Robinson Research Institute, School of Medicine, University of Adelaide, Adelaide, SA, Australia Robert T. Rydze Department of Obstetrics & Gynecology; Clinical Scientist Training Program, Baylor College of Medicine, Houston, TX, United States Alan Schneyer Department of Veterinary and Animal Science, University of Massachusetts-Amherst; Fairbanks Pharmaceuticals Inc, Concord, MA, United States Jeff Schwartz School of Medical Science, Griffith University, Gold Coast, QLD, Australia David B. Seifer Division of Reproductive Endocrinology & Infertility, Department of Obstetrics, Gynecology and Reproductive Sciences, Yale University School of Medicine, New Haven, CT, United States Dar-Bin Shieh Departments of Obstetrics and Gynaecology and Cellular and Molecular Medicine, Interdisciplinary School of Health Sciences, University of Ottawa; Chronic Disease Program, Ottawa Hospital Research Institute, Ottawa, ON, Canada; State Key Laboratory of Quality Research in Chinese Medicine, Macau Institute for Applied Research in Medicine and Health, Macau University of

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Science and Technology, Taipa, Macao; Institute of Basic Medical Science, Institute of Oral Medicine and Department of Stomatology, National Cheng Kung University Hospital, Advanced Optoelectronic Technology Center & Center for Micro/Nano Science & Technology, College of Medicine, National Cheng Kung University, Tainan, Taiwan; Interdisciplinary Program in Cancer Biology, Cancer Research Institute, and the Department of Obstetrics and Gynecology, Seoul National University College of Medicine, Seoul, Republic of Korea Ketan Shrestha Department of Animal Sciences, The Robert H. Smith Faculty of Agriculture, Food, and Environment, The Hebrew University of Jerusalem, Rehovot, Israel Joe Leigh Simpson Research and Global Programs, March of Dimes, White Plains, NY, United States Fred Sinowatz Department of Veterinary Sciences, Ludwig-Maximilian University Munich, Munich, Germany M.K. Skinner Center for Reproductive Biology, School of Biological Sciences, Washington State University, Pullman, WA, United States Yong Sang Song Departments of Obstetrics and Gynaecology and Cellular and Molecular Medicine, Interdisciplinary School of Health Sciences, University of Ottawa; Chronic Disease Program, Ottawa Hospital Research Institute, Ottawa, ON, Canada; State Key Laboratory of Quality Research in Chinese Medicine, Macau Institute for Applied Research in Medicine and Health, Macau University of Science and Technology, Taipa, Macao; Institute of Basic Medical Science, Institute of Oral Medicine and Department of Stomatology, National Cheng Kung University Hospital, Advanced Optoelectronic Technology Center & Center for Micro/ Nano Science & Technology, College of Medicine, National Cheng Kung University, Tainan, Taiwan; Interdisciplinary Program in Cancer Biology, Cancer Research Institute, and the Department of Obstetrics and Gynecology, Seoul National University College of Medicine, Seoul, Republic of Korea Jerome F. Strauss, III Department of Obstetrics and Gynecology, Virginia Commonwealth University, Richmond, VA, United States Qing-Yuan Sun State Key Laboratory of Stem Cell and Reproductive Biology, Institute of Zoology, Chinese Academy of Sciences, Beijing, China Yu Sun Key Lab of Stem Cell Biology, Institute of Health Sciences, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences and Shanghai Jiaotong University School of Medicine, Shanghai, China Reshef Tal Division of Reproductive Endocrinology & Infertility, Department of Obstetrics, Gynecology and Reproductive Sciences, Yale University School of Medicine, New Haven, CT, United States Enes Taylan Laboratory of Molecular Reproduction and Fertility Preservation, Department of Obstetrics, Gynecology and Reproductive Sciences, Yale University School of Medicine, New Haven, CT, United States

xiv

CONTRIBUTORS

Benjamin K. Tsang Departments of Obstetrics and Gynaecology and Cellular and Molecular Medicine, Interdisciplinary School of Health Sciences, University of Ottawa; Chronic Disease Program, Ottawa Hospital Research Institute, Ottawa, ON, Canada; State Key Laboratory of Quality Research in Chinese Medicine, Macau Institute for Applied Research in Medicine and Health, Macau University of Science and Technology, Taipa, Macao; Institute of Basic Medical Science, Institute of Oral Medicine and Department of Stomatology, National Cheng Kung University Hospital, Advanced Optoelectronic Technology Center & Center for Micro/Nano Science & Technology, College of Medicine, National Cheng Kung University, Tainan, Taiwan; Interdisciplinary Program in Cancer Biology, Cancer Research Institute, and the Department of Obstetrics and Gynecology, Seoul National University College of Medicine, Seoul, Republic of Korea

K.A. Walters School of Women’s & Children’s Health, University of New South Wales, Randwick, NSW, Australia

Chii-Ruey Tzeng Center for Reproductive Medicine, Department of Obstetrics and Gynecology, Taipei Medical University Hospital, Taipei, Taiwan

Han Zhao National Research Center for Assisted Reproductive Technology and Reproductive Genetics; Center for Reproductive Medicine, Shandong University; The Key Laboratory for Reproductive Endocrinology, Shandong University, Ministry of Education, Jinan; Center for Reproductive Medicine, Ren Ji Hospital, School of Medicine, Shanghai Jiao Tong University; Shanghai Key Laboratory for Assisted Reproduction and Reproductive Genetics, Shanghai, China

Barbara C. Vanderhyden Department of Cellular and Molecular Medicine, University of Ottawa; Cancer Therapeutics Program, Ottawa Hospital Research Institute; Department of Obstetrics and Gynecology, University of Ottawa, Ottawa, ON, Canada Maria M. Viveiros Department of Physiology and Pharmacology, College of Veterinary Medicine; Regenerative Biosciences Center (RBC), University of Georgia, Athens, GA, United States Ian N. Waldman Department of Obstetrics and Gynecology, Penn State College of Medicine, Hershey, PA, United States

Yun Wang Center for Reproductive Medicine, Department of Obstetrics and Gynecology, Peking University Third Hospital, Beijing, China Corrine Welt Division of Endocrinology, Metabolism and Diabetes, University of Utah, Salt Lake City, UT, United States Shannon N. Westin Department of Gynecologic Oncology and Reproductive Medicine, University of Texas MD Anderson Cancer Center, Houston, TX, United States Teresa K. Woodruff Northwestern University, Department of Obstetrics and Gynecology, Feinberg School of Medicine, Chicago, IL, United States Jie Yan Center for Reproductive Medicine, Department of Obstetrics and Gynecology, Peking University Third Hospital, Beijing, China

Xu Zhi Center for Reproductive Medicine, Department of Obstetrics and Gynecology, Peking University Third Hospital, Beijing, China Yi-Min Zhu Department of Reproductive Endocrinology, Women’s Hospital, School of Medicine, Zhejiang University, Hangzhou, China

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C H A P T E R

1 Follicle Selection in Mammalian Ovaries J.K. Findlay, K.R. Dunning, R.B. Gilchrist, K.J. Hutt, Darryl L. Russell, K.A. Walters Abbreviations AMH AR ArKO ARKO BMP15 CC COC DNA EGF ERα ERβ FSH FSHR GC GDF9 IGFI IGFII LH LHR MGC OSF NPPC NPR2 TGFα TGFβ ZP

new entity at the time of fertilization. It is critical, therefore, that this information is protected and nurtured during the life span of the mammal to ensure that a healthy oocyte is available for fertilization after ovulation during the fertile period. The mammalian ovary is the repository for all the oocytes, and it provides a specialized niche, the ovarian follicle, to protect and nurture the oocytes until they are selected for growth and ovulation, which can be up to 50 years in women.

anti-M€ ullerian hormone androgen receptor aromatase knockout mouse androgen receptor knockout mouse bone morphogenetic protein 15 cumulus cells cumulus-oocyte complex deoxyribonucleic acid epidermal growth factor estrogen receptor α estrogen receptor β follicle stimulating hormone follicle stimulating hormone receptor granulosa cells growth differentiation factor 9 insulin-like growth factor 1 insulin-like growth factor 2 luteinizing hormone luteinizing hormone receptor mural granulosa cells oocyte secreted factor natriuretic peptide precursor type C natriuretic peptide receptor 2 transforming growth factor α transforming growth factor β zona pellucida

The Ovarian Reserve By birth (human) or a few days thereafter (rodents), the ovary contains the maximum number of oocytes that a mammal will have for the rest of its life. This pool of primordial follicles is called the ovarian reserve. Thereafter, the size of the pool decreases at variable rates until such time as the ovary is devoid of oocytes, defined as menopause (human) or infertility (rodents). The number of oocytes at birth and their rate of decline determine the fertile life span of the mammal. There is no compelling evidence in neonates or adults that under physiological circumstances, new oocytes are formed once the initial pool is established.

This chapter focusses on the factors and molecular mechanisms responsible for the selection for growth or death of follicles during all stages of folliculogenesis. It discusses (a) selection of primordial follicles in the ovarian reserve for activation or death prior to puberty and during the fertile life span, (b) the intra- and extragonadal control of selection of antral and ovulatory follicles, and (c) oocyte regulation of folliculogenesis through autoand paracrine interactions within the follicle, including mechanisms responsible for polyovulation.

Mammalian Follicles and Folliculogenesis Ovarian follicles each containing an oocyte are defined as either nongrowing or growing (Fig. 1). Nongrowing follicles are primordial follicles, which are characterized by an oocyte surrounded by squamous or flattened granulosa cells (GCs). The DNA in the oocytes of these follicles is arrested in meiotic prophase I and will remain so until ovulation. Only primordial follicles make up the ovarian reserve and have been described variously as “dormant” or “resting,” although oocytes in these primordial follicles have an active metabolic profile and the DNA damage repair machinery to guard the integrity of the genome (see below). The appearance of cuboidal GC with or

INTRODUCTION The mammalian oocyte or germ cell contains all the genetic information that the female will transfer to the The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00001-7

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1. FOLLICLE SELECTION IN MAMMALIAN OVARIES

Gonadotropin-dependent growth antral Gonadotropin-independent growth preantral

Antral/preovulatory (dominant) Early antral

Secondary Activated Primordial primordial

Oocyte Granulosa cell

Primary

Zona pellucida

Basement membrane

Theca cells

Antrum Granulosa cells

Cumulus cells

Atresia

FIG. 1 A schematic representation of the stages of mammalian folliculogenesis, follicle types and highlighting atresia, and gonadotropin dependence.

without enlargement of the oocyte is morphological evidence of activation of primordial follicles to enter an early growing phase. Primary follicles are formed after complete transformation of the squamous GC to cuboidal, a process thought to be initiated by a collaboration between the oocyte and the GC. Ultimately, resolution of what constitutes selection and activation of primordial follicles will need identification of the molecular pathways responsible for the changes in oocyte and GC structure and function. Current knowledge of this topic is reviewed below. Early follicle growth consists of enlargement and proliferation of GC, enlargement of the oocyte, formation of a basement membrane surrounding the GC, and formation of a zona pellucida (ZP) surrounding the oocyte (Fig. 1). Based on these morphological characteristics, follicles in the early growth phase are defined as preantral or primary and secondary follicles (Fig. 1) in which there are one or more layers of GC surrounded by a basement membrane and the ZP around the oocyte. Formation of a theca layer and an antrum transition follicles to the antral stage. Differentiation of the GC into cumulus cells (CCs) surrounding the oocyte and mural GC lining the basement membrane, and expansion of the antrum readies the follicle for ovulation. Death of follicles occurs at all stages of folliculogenesis. As puberty approaches, the ovary acquires the full complement of different follicle types at all stages—primordial, growing (preantral and antral), and atretic. But ovulation does not occur until after puberty. The time a follicle takes to develop from the primordial stage to ovulation is up to 12 months in women and approximately 3 and 4 weeks in rodents. The largest portion of that time is taken up by preantral

folliculogenesis, whereas the later antral stages that undergo cyclical maturation in waves take only days and are regulated by the hormonal profiles and intrafollicular milieu of growth factors during the follicular phases of the menstrual or estrous cycle. Details of many of the molecular pathways in the oocyte, those responsible for the differentiation and proliferation of the different somatic cell types that make up growing follicles, and how these underpin the interactions between the oocyte and its somatic cell niche, are now understood and are reviewed below. Of particular importance has been the establishment that (a), growth of preantral follicles can be independent of gonadotropins, particularly FSH, whereas antral follicle growth is FSH and luteinizing hormone (LH) dependent and (b) the presence of a healthy oocyte, acting in an autocrine and paracrine fashion with the somatic cells in the follicular niche, is essential for the ultimate fate of the follicle (see below). Furthermore, the influence of the oocyte specific factors may underlie the difference between single and multiple ovulations.

Selection of Ovarian Follicles The focus of this chapter is the selection of follicles during all stages of folliculogenesis. The primordial follicle pool or ovarian reserve decreases with age for two reasons: the primordial follicles either die mostly by atresia or are selected for growth and enter the pool of growing follicles, throughout which most die, but a select few are ovulated and gain the chance to be fertilized and form a new organism. The location and properties that

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determine the choice of primordial follicles to be selected from the ovarian reserve for activation are not understood. The focal point of atresia in primordial follicles is the oocyte, and recent research has thrown new light on the role of the intrinsic apoptotic pathways, particularly the BH3-only factors, as opposed to the DNA repair pathways guarding the genome (see below). In growing follicles, the focal point of atresia is the GC. Selection of those antral follicles that enter the final growth phases ultimately to become dominant Graafian follicles that ovulate is based on their capacity to avoid atresia due to protection by locally produced growth factors and steroids, enabling them to survive reduced FSH stimulation in the dominant phase of growth immediately preceding ovulation. Furthermore, Graafian follicles have acquired the molecular machinery to produce sufficient estradiol17β to trigger the ovulatory surge of LH with consequent ovulatory changes in the oocyte and follicular somatic cells. The oocyte also influences the selection processes by providing checkpoints during folliculogenesis, particularly after antrum formation. In addition to the important intrafollicular role played by the somatic cells of the follicle to nurture and interact with the oocyte, these cells mainly in antral follicles are also essential for production of the sex steroid hormones (estradiol-17β, progesterone), which are responsible for the secondary sex characteristics of the female and to prepare the uterus for pregnancy, and provide substrate (androgens) for the production of estradiol. These somatic cells also produce peptides (e.g., inhibin) which together with the sex steroids facilitate the negative and positive feedback effects of the ovary on the hypothalamic–pituitary axis to regulate the levels of gonadotropin during the estrous or menstrual cycles.

SELECTION OF PRIMORDIAL FOLLICLES The ovarian reserve of primordial follicles is established early in life and it is from this stockpile that all growing follicles and ovulated oocytes are derived. Because of this, the number of primordial follicles initially formed in the ovary, and the rate at which they are activated to begin further development, or triggered to die, greatly influences the female fertile life span. Of note is the observation that in all mammalian species analyzed, there are many more primordial follicles than required for ovulation. For example, in humans 1 million primordial follicles are present at birth and this number is reduced to around 300,000 at puberty, with approximately 1000 remaining when the menopause begins [1,2]. During the time between puberty and end of the reproductive life span, only 400–500 human oocytes will be ovulated. Why an excess of primordial

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follicles is formed is not known. However, it is clear that the selection of an individual primordial follicle for activation, vs continued quiescence or death, represents the first point of selection in the step-wise process of folliculogenesis that ultimately culminates in the production of a meiotically mature and developmentally competent oocyte (Fig. 1).

Selection for Death Shortly after the ovarian reserve is established, a large number of primordial follicles are eliminated from the ovary as part of normal, developmentally regulated processes. Using empirical data and mathematical modeling, Tingen and colleagues estimated that approximately 155 primordial follicles per ovary per day undergo atresia in young mice (postnatal days 6–19) [3]. Given that approximately half this number was assumed to have been activated, it was concluded that death of primordial follicles contributes to greater depletion of the ovarian reserve than activation during the early stages of life [3]. There are a number of possible explanations for the elimination of primordial follicles in the days immediately following their formation. For example, follicles containing lowquality oocytes (e.g., those with damaged genomes or insufficient organelles) may be actively eliminated [4]. It is also possible that primordial follicles with inadequate numbers of GC are selected for death [5]. A passive trigger for primordial follicle death may be inadequate growth factor support [5]. Interestingly, the onset of puberty has also recently been identified as a critical developmental window during which primordial follicles are lost in large numbers. Liew and colleagues showed that 50% of primordial follicles present in the mouse ovary are actively eliminated at puberty and that this process is triggered by gonadotropins and mediated by the pro-apoptotic protein, BMF [6]. Why such a large cohort of primordial follicles are selected for death is not known, but one hypothesis is that a quality control checkpoint exists at puberty to ensure that only those follicles of the highest quality are available for adult reproduction. In addition to the normal loss of primordial follicles due to physiologically regulated processes, exogenous factors, such as exposure to anticancer treatments and environmental toxicants, can accelerate primordial follicle loss [7–10]. In this instance, primordial follicles are actively selected for elimination by apoptosis because they have sustained damage that could potentially compromise their ability to give rise to healthy offspring after fertilization [10]. This may be damage to the genome, or vital organelles such as mitochondria. In particular, it is essential that the oocyte genome is under rigorous surveillance, to ensure that DNA damage is detected and repaired, and that those oocytes with DNA damage that

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cannot be repaired are eliminated. These processes are critical to protect against the introduction of germline mutations. In this regard, the transcription factor TAp63, a p53 family member, is a key quality control factor for primordial follicles and has been referred to as the guardian of germ line [11,12]. The TAp63 isoform is expressed at high levels in primordial follicle oocytes and it is an essential mediator of DNA damage-induced oocyte death through transcriptional activation of the proapoptotic Bcl-2 family member PUMA [10].

Selection to Activate The activation of primordial follicles is characterized at the morphological level by growth of the oocyte and proliferation and differentiation of the surrounding GC. Once this process is initiated, it cannot be reversed. Because the commitment to further development is unidirectional, the rate of primordial follicle activation must be tightly controlled to ensure that sufficient reserves remain in order to support fertility throughout reproductive life [13]. Indeed, the most common mouse models of premature ovarian failure are those in which primordial follicles

become activated in an uncontrolled manner, such that the ovarian reserve is prematurely depleted [14]. In recent years, significant progress has been made toward characterizing the molecular pathways that govern the transition of follicles from a state of dormancy to growth. In particular, studies in mice have revealed the importance of the PI3K/PTEN-Akt-FOXO3 cascade in controlling oocyte growth associated with primordial follicle activation [15–20] (Fig. 2). Oocyte-specific deletion of Tsc1 and Tsc2 in mice results in global recruitment of primordial follicles, implicating mTORC1 signaling in the control of primordial follicle activation [21]. In addition, oocyte-specific transcription factors Lhx8, Sohl1, and Nobox are also essential for maintaining dormancy or regulating activation [22–24]. Despite an improved understanding of the complex signaling networks responsible for mediating the activation process, the question remains, “What triggers the pathways that result in activation in some follicles but not others?” In this regard, growth factors produced within the ovarian microenvironment are thought to be key regulators of follicle activation. In particular, the production of the growth factor Kit Ligand (also known as stem cell factor) by GC is an important pro-activation

FIG. 2 The PI3K signaling pathway in the oocyte of primordial follicles. Reproduced from Adhikari D, Liu K. Molecular mechanisms underlying the activation of mammalian primordial follicles. Endocr Rev 2009;30(5):438–64, by permission Oxford University Press.

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SELECTION OF PRIMORDIAL FOLLICLES

signal [25,26]. Kit Ligand likely exerts this effect by binding and activating KIT at the oolema, which in turn signals through PI3K [26]. In addition to the requirement for activators, primordial follicles are retained in a quiescent state by locally produced inhibitory factors. For example, AMH and activin production by growing follicles have roles in suppressing the rate of activation [27,28]. Primordial follicles were less likely to grow when a neighboring primordial follicle was in close proximity [29], suggesting that, similar to growing follicles, primordial follicles secrete a diffusible factor that prevents nearby primordial follicles from activating. Such an inhibitory molecule would be particularly important for controlling the rate of follicle activation prior to the first wave of folliculogenesis, when growing antral follicles are not yet present. Thus, in addition to the role of specific growth/inhibitory factors, the spatial relationships between follicles, potentially creating morphogen gradients in the ovary, may play an important role in the selection of individual primordial follicles for activation. In 1968, Henderson and Edwards proposed the production-line hypothesis to explain the differential selection of follicles for activation and increased trisomy rate in the offspring of older women [30]. This theory posits that the first oocytes to enter the meiotic prophase during the initial establishment of the ovarian reserve are also the first selected to activate and subsequently ovulate. Secondly, it states that those oocytes that enter meiosis first also have the highest crossover frequency, and thus are less prone to meiotic nondisjunction, thereby implying that “meiotic quality” is a selection factor for activation. This hypothesis was formulated on the basis of studies showing that chiasmata frequency is associated with the timing of meiotic entry in mice. However, a subsequent study in humans could not find any evidence for decreased crossover frequency between oocytes entering meiosis late in fetal development compared to those entering meiosis at earlier time point [31]. Even though the timing of meiotic entry did not correlate with chiasmata number in oocytes, this study does not rule out the possibility that in humans primordial follicles containing oocytes with high crossover numbers activate first, such that in women of advanced maternal age only oocytes with few chiasmata remain. Interestingly, a small study of IVF-treated women suggested that risk of a trisomic pregnancy is increased when the ovarian reserve is small, and this risk is independent of age [32]. The most simplistic interpretation of these data is that highest quality primordial follicles are selected to activate first and when the supply of healthy primordial follicles becomes reduced, regardless of age, the availability of high-quality oocytes is also reduced. The relationship between the location of follicles in the ovary and the order of activation has also received considerable attention. In separate studies, Hirshfield and

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Byskov reported that the first follicles to activate after the ovarian reserve is established are those located in the central medulla region of the ovary [33,34]. Recent works extend these earlier observations and propose that there may be two pools of primordial that are distinct in the timing of their formation, their location within the ovary, and their functional roles [35,36]. During endowment of the fetal mouse ovary, the primordial follicles that assemble first are located in the central region of the ovary and these are also the first cohort to activate. Lineage tracing studies in mice showed that most of the primordial follicles making up this first group are used before postnatal day 60 and thus primarily contribute to the establishment of sexual maturity and endocrine cyclicity during early life [35,36]. The second population of primordial follicles is formed in the ovary slightly later, is located in the cortex, and is primarily responsible for supplying follicles to support adult fertility [35,36]. That these two pools (i.e., comprising follicles that activate prevs postpubertally) are functionally different is supported by studies in cattle showing that oocytes retrieved from prepubertal animals and matured in vitro are developmentally poor compared to oocytes isolated from adults [37]. Furthermore, Anderson and colleagues reported the presence of unhealthy primordial follicles, characterized by abnormally large oocytes, in the ovarian cortex of prepubertal girls, which could not be detected in the adult [38]. These data suggest a selective process may exist in which primordial follicles containing low-quality oocytes are preferentially activated and/or eliminated from the ovary prior to the onset of adult reproductive capacity.

Dynamic Changes in the Selection for Death or Activation of Primordial Follicles Over the Fertile Life Span Studies suggest that the rate at which primordial follicles are selected for activation and/or elimination is dynamically regulated and varies from birth to the end of the fertile period. Indeed it appears the number of follicles that activate is greatest early in life and then reduces in adulthood [39]. In line with this idea, animal studies suggest that the number of follicles that activate is proportional to the total size of the ovarian reserve. If this is true then one would expect greater absolute numbers of growing follicles to be present early in life, with lower numbers later in life. Studies in cattle [40] and mice [41] indicate that a large ovarian reserve of primordial follicles in young females is associated with larger numbers of growing follicle than observed in older females, which have smaller reserves of primordial follicles. In some cases, while the absolute number of follicles that activate changes with age, the proportions of follicles making up the quiescent vs growing fractions remain fairly constant

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[41]. However, it is important to note that there is conflicting evidence regarding the rate of follicle activation with increasing age in humans. Some studies suggest that the rate of activation increases in the years preceding menopause [42–46]. However, a more recent model of human follicular dynamics, based on a larger histological dataset, suggests that the rate of primordial follicle activation increases from birth to 14 years and then declines until menopause [2]. This later model predicts that in women with an average age of 49 years at menopause, a maximum of 900 primordial follicles are recruited each month at 14 years of age, while at 40 years of age, only 100 primordial follicles are recruited each month [2]. Thus, fewer follicles activate and make up the growing follicle population in older women, likely due to a proportionally smaller reserve. There is also evidence to suggest that the rate of primordial follicle activation is dynamically regulated not only in response to physiological depletion of the ovarian reserve associated with age but also due to pathological depletion. This concept is supported by studies in which the ovarian reserve is significantly depleted by exposure to γ-irradiation, for example, but the loss of fertility is only modestly affected [9]. Thus, in this context of a dramatically reduced ovarian reserve, it is possible that the rate of activation is reduced in order to sustain fertility over an adequate period of time. How these changes in the selection of primordial follicles for activation might be mediated is not known but represent an exciting avenue for future research.

INTRA- AND EXTRAGONADAL HORMONAL CONTROL OF FOLLICLE SELECTION As described earlier, a cohort of primordial follicles is activated at regular intervals forming a pool of growing follicles that eventually reach the antral stage (Fig. 1). Rising FSH concentration during the follicular phase of the cycle induces the emergence of a follicle wave from among this pool of growing antral follicles (Fig. 3). In most large mammals, including humans, only one follicle is intended to ovulate and release the oocyte into the fallopian tube for fertilization. The remainder of this growing follicle pool regress through atresia. Of all the oocytes present in the ovaries at birth, only 0.1% will actually be ovulated. The process by which the follicle that will ovulate is selected to become dominant is another facet of follicle selection. The reason for this relatively inefficient method of producing a single dominant follicle is not clearly understood but studies of follicle selection in several species show it is a complex and responsive process whereby the molecular interaction of the follicle/oocyte to the

hormonal milieu critically determines the “successful” follicle. Tracking of the growth rates of follicles by ultrasound imaging in cows and horses indicates that the follicles competing to achieve dominance fluctuate in their rates of growth and relative size in the preselection period until dominance is eventually exerted by one follicle and the growth of subordinate follicles slows and they subsequently regress [47,48] (Fig. 3). In experiments where the large putative dominant follicles are ablated, the smaller subordinate follicles will increase their growth rates and fill the void, with one of them arising to become selected as the dominant follicle [49]. These findings illustrate that there is a competitive aspect to follicle selection with the larger follicles “subordinating” the remainder of the cohort by suppressing their growth and development. These characteristics of the growth and selection process support the hypothesis that it may, in part, fulfill a quality requirement whereby the follicle/oocyte unit with the most robust capacity to respond to the cyclic fluctuations in endocrine and paracrine environment achieves a growth advantage which is reflective of the “quality” of the oocyte. The capacity of the oocyte to influence selection by promoting proliferation and suppressing apoptosis in follicular somatic cells is reviewed below. The expression of dominance involves serial physical and biochemical changes among the follicles in the growing cohort, which eventually leads to one selected follicle. The growing follicles all produce endocrine and paracrine acting factors, which are drivers of the female hormonal cycle. Early after their activation, follicles grow mainly under control of paracrine factors and independent of FSH. Steroids, inhibin, and AMH are important regulators of growth or atresia [50]. The transition to antral follicles is FSH dependent [51,52], and antral follicle growth requires FSH support. As deviation of the dominant follicle emerges, their growth becomes less dependent on FSH and they secrete FSH-suppressing factors inhibin and estradiol, reducing circulating FSH and removing the tropic support that subordinate follicles require [53].

Gonadotropin-Independent Follicle Growth— Prenatal to Antral Development The early preantral and the preantral to early antral transition stages of follicle development can occur independently of extraovarian gonadotropin support. Mice genetically lacking gonadotropins exhibit follicles at the early stages of development, but mice deficient for β-FSH or the FSH receptor (FSHR) gene exhibit follicular developmental arrest at the preantral stage [51,52], and LH receptor (LHR)-knockout mice exhibit a block in follicle development at the antral stage [54]. Likewise FSH is essential for progression to the antral stage in humans as

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20

1st wave anovlatory

9

Dominance

2nd wave anovlatory

3rd wave ovulatory

1st wave etc...

Dominance Ovulation

5

Follicle diameter (mm)

Selection Follicles recruited for continued growth

3

Selection

Selection

Recruitment Recruitment

0.5

Atresia

Cohort of initiated follicles

Atretic follicles

0.2

Primordial 0.04

Depletion Initiation

Early follicle growth

Follicle selection

Approximately 4 months Early follicle development

0

7

14

21/0

7

Days of oestrus cycle

FIG. 3

Patterns of follicular growth in the bovine. Follicle growth is a lengthy process with the majority of time spent in the early stages of development, while the terminal stages occur in a wave-like pattern. Follicle growth occurs in a cascade-like fashion with cohorts of primordial follicles being triggered to grow; however, most are destined to undergo atresia. Growth of committed follicles to the gonadotropin responsive stage is approximately linear, hence ensuring a continuous supply of gonadotropin-responsive follicles. A rise in FSH leads to the emergence of a cohort of gonadotropin-dependent follicles, and from this group one or a group of potentially ovulatory follicles arise. In the presence of a corpus luteum, a nonovulatory wave occurs and the dominant follicles undergo atresia. However, if the corpus luteum has regressed, an ovulatory wave occurs and the dominant follicle ovulates, releasing the oocyte.

evidenced by the lack of antral follicles in hypogonadal individuals. While LH and FSH are not essential for early stages of growth, there is evidence to support a beneficial effect for them during the gonadotropin-independent stage. FSHR is present on the GC from the early preantral follicle stage [55] inferring a functional role of direct FSH actions. In addition, follicles did not progress beyond the two cell layer stage in human ovarian xenografts in mice homozygous for severe combined immunodeficiency (SCID) and hypogonadism (low FSH). However in FSHtreated grafts, follicles were shown to develop to the antral stages [56]. LHR are expressed on theca cells from preantral and antral follicles, and studies have suggested that LH is a survival and differentiation factor that facilitates oocyte development in the presence of FSH [57]. These findings infer that while gonadotropins are not essential for the early stages of follicle growth, they may have a beneficial effect on the early follicular growth phase. Gonadotropins have a close interrelationship with steroids. The secretion of FSH and LH from the pituitary

is mainly regulated by the steroids (and inhibin in the case of FSH) secreted from the gonads, and FSH and LH themselves stimulate steroid synthesis in the process termed the two-cell two gonadotropin theory of estrogen biosynthesis [58]. Within the follicle the GC and theca cells make up the follicular unit that synthesizes steroid hormones. Thecal cells are able to bind LH, which stimulates the production of androgens from cholesterol. These androgens diffuse into the neighboring GC where they can modulate cell growth and survival as well as be converted into estrogens by the FSH-induced activation of the enzyme aromatase [58]. Estrogen production occurs from the secondary stage and increases dramatically at around the time of antrum formation when GC, distinct from the CC phenotype, emerge and these GC are highly sensitive to FSH. The ovaries are the primary source of estrogens, which play a fundamental role in the development of a follicle, highlighted by the fact that mice deficient for estrogen receptor genes (ERα and ERβ) display decreased or a

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complete loss of fertility [59–61]. Furthermore, the aromatase-knockout (ArKO) mouse, which is unable to make endogenous estrogens due to the loss of the enzyme aromatase, is infertile with follicles arrested at the antral stage [62]. Estradiol, the principle estrogen, has a wellestablished role in the feedback regulation of LH and FSH secretion evident by the marked increase in circulating FSH and LH levels in ArKO mice [62]. Within the ovary both ER isoforms are present from the primary follicular stage [63]. An increase in levels of estradiol in the follicular fluid is one of the earliest signs of selection of a follicle to achieve dominance [64]. Along with FSH and insulin-like growth factor-I (IGF-I), estradiol stimulates the proliferation and differentiation of GCs [65]. ERα is required for follicle dominance as female mice with a loss of ERα display a block in follicle development at the antral stage [61], while mice with a deletion of the ERβ gene display a less compromised phenotype, with some exhibiting follicle arrest but others displaying follicles at all stages but reduced numbers of corpora lutea [60]. These results imply that estradiol can exert varying effects via its different receptors, with ERα proposed to facilitate the proliferative actions of estradiol, while ERβ is thought to mediate the differentiative effect of estradiol. Apart from the role of androgens in ovarian function as the obligatory precursors for estrogen biosynthesis, it is now evident that they also play a signaling role in regulating follicle development. The development of androgen-resistant female mouse models (ARKO) has confirmed that while androgen actions are not required for full folliculogenesis to occur, they play an important role in optimizing ovarian function. ARKO mice are subfertile and display dysfunctional follicle development, increased follicular atresia, and reduced ovarian expression of key regulators of follicle health, including FSHR and IGF1 receptors [66–68]. The androgen receptor (AR) is expressed throughout most stages of follicle development, with expression found to be most abundant during the early stages. Evidence overall supports a stimulatory role for androgens in the early follicular developmental phase [69]. In vitro culture of mouse preantral follicles with different androgens (testosterone, androstenedione, dehydroepiandrosterone, and dihydrotestosterone) was shown to enhance follicle growth and development [70,71], and importantly, these stimulatory effects were blocked by an AR antagonist [70]. Similarly, both bioactive androgens, testosterone and dihydrotestosterone, increased the number of preantral and small antral follicles in primate ovaries [72]. A synergistic interaction between androgens and FSH appears to be important as treatment of primates [73] and mice [74] with androgens increased FSHR expression and enhanced FSH-mediated preantral to antral follicle development in mice [74]. Furthermore, mouse preantral follicle responsiveness is improved by testosterone [71].

While it is now clear that androgens exert important positive effects on early follicular growth and health, there is an emerging theme that an appropriate balance in androgen actions is key for the maintenance of optimal ovarian function. Rodent, sheep, and primate animal models have highlighted that abnormally elevated androgen levels disrupt the crucial balance required for normal follicular development and can induce arrest of follicle growth, which is characteristic of polycystic ovary syndrome in women [75–77].

Gonadotropin-Dependent Follicle Growth and Preovulatory Follicle Selection During the antral stage of development, the most advanced cohort of gonadotropin-responsive follicles emerge concomitantly with the increase in FSH, to form the pool of gonadotropin-dependent follicles. Rising FSH levels in the early follicular phase of the cycle stimulates a wave of follicle growth and differentiation from which the dominant follicle(s) will subsequently emerge [78,79]. Increasing pulsatile release of LH also promotes follicle development as well as steroidogenesis [80]. These follicles grow and regress in a regular sequential pattern of waves largely governed by the secretion of FSH, IGF1, estradiol, and inhibin (Fig. 3). At this follicular stage, gonadotropins are essential for development, and the promotion by FSH of estradiol synthesis by the GC is of key importance. Estradiol production increases dramatically and is maximal in the dominant follicle prior to ovulation. The actions of FSH to enhance aromatase activity are increased through the actions of IGF-I, IGFII, and estradiol itself, and conversely, are inhibited by follistatin and EGF or TGF-α [81]. Estradiol in the follicle has a paracrine role promoting GC proliferation through induction of cyclin D2 [82] and thus follicle growth. The sustained increase in circulating estradiol and inhibin levels at this stage has an endocrine role acting on the pituitary to reduce FSH gene transcription and protein secretion leading to a gradual decline in FSH acting on the growing follicle pool. Progressively rising estradiol is also the endocrine trigger that acts on the hypothalamus to increase GnRH secretion and on the pituitary to increase GnRH sensitivity leading to the ovulatory LH surge [83–85]. Hence, estradiol has many key actions in directly regulating gonadotropin secretion as it switches from suppressing levels through negative feedback during the early stages of follicle development to exerting a stimulatory effect during the terminal follicular stages via a positive feedback mechanism. At this stage of folliculogenesis, deviation occurs with one dominant follicle selected in monovular species or a cohort of dominant follicles in polyovular species [53]. As described later, these differences in ovulation rate could be due to the influence of growth factors produced by

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the oocyte. The mechanism of dominant follicle selection has been most studied in large monovular species (horses and cows) where follicle growth can be tracked by ultrasound imaging and follicular fluid and ovarian cells can be sampled from individual follicles with known growth dynamics [53]. Importantly the dominant follicle is slightly larger at the time it is selected, but also develops a more robust vascular bed and higher blood flow [86]. Indeed, assessment of vascular flow in human follicles using Doppler ultrasound at the time of oocyte pickup during assisted reproduction has been found to be a good surrogate indicator of the oocyte with highest developmental potential [87]. Higher expression of LHR and FSHR enhance the dominant follicle sensitivity to gonadotropins [53]. Concentrations of factors important for vascular development and proliferation, VEGF, IGF1, and estradiol, become higher in the follicular fluid of the dominant follicle. IGF can be inhibited by IGF-binding proteins [88,89], hence low levels of these are also necessary for the accelerated growth in dominant follicles [90]. IGF1 and estradiol are potent mitogenic factors in GC. The IGF system is particularly important in the follicle selection process as IGF-I has been shown to stimulate steroidogenesis in theca cells and proliferation, differentiation and secretion of estradiol in GC. Furthermore, IGF-I increases the sensitivity of small follicles to gonadotropin stimulation and hence push forward their growth even in the phase of declining FSH which deprives subordinate follicles of necessary growth support [91]. Two forms of inhibin (A and B) are secreted by follicles and both can repress FSH secretion, but inhibin B may be the more important inhibin isoform in expression of dominance in the human ovary due to its higher levels and tighter inverse association with FSH circulating profile [92]. Inhibin reduces the circulating concentration of FSH and acts in the follicle to increase androgen production [93]. An intrafollicular role of inhibin in promoting selection of follicles is also evident. Immunoneutralization of inhibin has been convincingly shown to increase the number of ovulating follicles [94] with high embryo development outcomes [95]. In part this is through reducing feedback inhibition of FSH secretion, but the increase in ovulation cannot be replicated with high FSH treatment alone and a marked synergistic effect of inhibin antiserum plus equine chorionic gonadotropin treatment indicates that inhibin neutralization has unique effects on ovulation rate, and inhibin antiserum has been shown to directly affect gene expression in GCs [96]. Other studies showed that neutralization of the N-terminal peptide of inhibin-α also increased circulating FSH and the number of preovulatory follicles in sheep, yet the follicles failed to ovulate [97] further suggesting that inhibin influences follicle selection to ovulate through both endocrine and intraovarian pathways. Apart from their role as the substrate for estrogen production, androgens have less of an influence on the later

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stages of follicle development. Treatment of pigs with testosterone or dihydrotestosterone during the late follicular phase increased preovulatory follicle numbers [98,99]. However species differences exist, as testosterone and dihydrotestosterone have no effect on primate preovulatory follicle numbers [72]. Evidence also implies a direct role for androgens in oocyte maturation as testosterone promotes in vitro germinal vesicle breakdown in murine oocytes, which is suppressed by the addition of an AR blocker [100].

OOCYTE REGULATION OF FOLLICLE SELECTION Oocytes Participate in Follicle Selection via Key Paracrine Messengers One of the most important new concepts to emerge over the course of the past one to two decades in relation to follicle selection in mammals is the role the oocyte itself plays in this process. The traditional perspective of the relationship between the follicle and its oocyte is that the oocyte’s growth and development is determined by the follicle and that the oocyte is passive in terms of the regulation of folliculogenesis. We now know that the oocyte plays a major role in controlling folliculogenesis, including, promoting early follicle growth, determining the species-specific ovulation quota, controlling granulosa cell lineage differentiation affecting oocyte development and quality, and establishing the capacity for ovulation [101]. The oocyte achieves these actions principally through the production and paracrine actions of two growth factors: growth differentiation factor 9 (GDF9) and bone morphogenetic protein 15 (BMP15). These TGFβ superfamily members are most closely related to each other and there are notable interactions (in many cases synergies) between GDF9 and BMP15, which are likely mediated by the GDF9:BMP15 heterodimer, cumulin. These growth factors are also notable in that they are generally regarded as gamete-specific factors, making them potentially attractive therapeutic and diagnostic targets in human and veterinary reproductive medicine.

GDF9 and BMP15 Regulate Follicle Development, Selection, and Ovulation Rate The capacity of the oocyte to control follicle development and ovulation rate is mediated by secretion of the paracrine factors, GDF9 and BMP15, which act on GC throughout folliculogenesis. The essential role of BMP15 and GDF9 in folliculogenesis and female fertility is evidenced by the numerous studies utilizing gene ablation in mice, naturally occurring mutations in sheep and humans, immuno-neutralization in ruminants, and

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in vitro studies employing recombinant proteins. Collectively, these studies show that oocyte control of GC function is a critical requirement for committed follicles to transition to the FSH-responsive and FSH-dependent stages in folliculogenesis [53]. Oocyte secretion of GDF9 is essential for follicle development and fertility in female mammals. GDF9 has been detected in the oocytes of mice, rats, cows, sheep, goats, pigs, rhesus monkeys, and humans [102]. GDF9 is found exclusively in oocytes from the primary stage onward in human, mice, and rats, and in sheep and cow oocytes beginning at the time of primordial follicle formation [102]. The essential role of the oocyte in early folliculogenesis was first demonstrated with the generation of the Gdf9-knockout mouse. The female Gdf9 null mouse is sterile due to a block at the primary to secondary stage of follicle development demonstrating the requirement for GDF9 for the growth of primary follicles in the mouse [103]. Similarly, female sheep with homozygous naturally occurring mutations in GDF9 are sterile due to a block in follicle development at the same stage as the Gdf9 null mouse [104]. These observations are supported by in vitro studies whereby recombinant GDF9 promotes follicle growth to the secondary stage [105] and growth of secondary follicles [106]. The requirement of GDF9 for the transition from the primary to secondary stage is partly attributable to the potent proliferative effects of GDF9 on primary GC [107]. The block in follicle development observed in the Gdf9-knockout mouse may also be due to dysregulation of inhibin-α as folliculogenesis in mice null for both Gdf9 and Inha proceed beyond the primary stage [108]. In addition to the effects of GDF9 on GC, it appears that GDF9 is also involved in the recruitment of theca cells from the ovarian stroma in mice and theca cell differentiation and function in human, mouse, and rat [102]. In contrast to GDF9, the requirement of BMP15 for fertility differs markedly between species. In broad terms, BMP15 is required and has important regulatory effects in monovular mammals, whereas it has a negligible role in polyovular mammals. BMP15 remains undetected in sheep and mouse oocytes until the primary stage [102]. In the mouse, although oocytes express Bmp15 mRNA throughout most of folliculogenesis, production of mature and active BMP15 is thought to be minimal until the ovulatory LH-surge when it is upregulated [109,110]. Consistent with this, the phenotype of the Bmp15 null mouse is in stark contrast to the Gdf9 knockout mouse. Female Bmp15 null mice are mildly subfertile and display no abnormalities in early follicle development [111]. Thus, LH-induced processing of BMP15 in polyovular species likely limits functionality of BMP15 to oocyte maturation, cumulus expansion, and ovulation as evidenced by the phenotype of the Bmp15 null mouse. Conversely, in monovular species, such as sheep and humans,

processed and active BMP15 is thought to be present in the oocyte from the primary stage onward, and BMP15, in conjunction with GDF9, is regarded as central in follicle development and selection. Naturally occurring inactivating mutations in BMP15 result in sterility in sheep due to a failure of folliculogenesis to proceed beyond the primary stage [104,112]. Interestingly, ewes heterozygous for either BMP15 or GDF9 inactivating mutations have an increased ovulation rate and fecundity. The effects of mutations in BMP15 and GDF9 on fertility in sheep have been replicated in studies where ewes were immunized against these proteins. Complete immunoneutralization results in a block at the primary stage of follicle development while partial neutralization leads to increased rates of ovulation and fecundity [113]. The mechanism by which partial actions of GDF9/BMP15 (whether by gene dosage or immunization) result in an increase in ovulation rate, but inaction of these factors induces sterility, has intrigued reproductive biologists for some time. GDF9 and BMP15 are potent regulators of GC-secreted inhibin and steroids and control gonadotropin receptor responsiveness. However, curiously, highly fecund ewes that are heterozygous for the inactivating mutations in GDF9 or BMP15 have normal endocrine levels of FSH, LH, estradiol, progesterone, and inhibin and normal primordial follicle numbers [114]. It appears the mechanism underlying the increased fecundity in animals with altered GDF9/BMP15 function is the premature acquisition of functional LHR in smaller and in more follicles, which hence ovulate more follicles at a smaller size, than in wild-type animals (Fig. 4) [114,114a]. Hence, GDF9 and BMP15, secreted by the oocyte, control ovulation rate and fecundity principally by regulating GC receptivity to gonadotropins. These intriguing findings in domestic animals inspired a search for GDF9 and BMP15 mutations affecting human fertility. Numerous heterozygous missense mutations in BMP15 have been identified in women with premature ovarian failure [115]. The differing mutations lie principally in the prodomain of BMP15, affecting the production and activity of BMP15 proteins and also its capacity to interact with GDF9 [116]. Aberrant BMP15 and/or GDF9 function are also implicated in human dizygotic twinning, the ovarian defect in Turner syndrome and ovarian reserve, polycystic ovarian syndrome and particular patient haplotypes associated with responsiveness to superovulation [115]. In monovular mammals, GDF9 and BMP15 are coexpressed in the oocyte for nearly all of folliculogenesis, and as they are closely related and are known to interact with each other, they are usually thought of as acting in combination in these species. This perspective was initially motivated by the observation that ewes carrying heterozygous mutations in both GDF9 and BMP15 have

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FIG. 4 Altered BMP15 activity changes the timing of LH receptor acquisition leading to polyovulation. Reproduced from McNatty KP, Juengel JL, Pitman JL. Oocyte-somatic cell interactions and ovulation rate: effects of oocyte quality and embryo yield. Reprod Biol Insights 2014;7:1–8, with permission.

even higher fecundity than animals with mutations in either GDF9 or BMP15 alone [104]. This genetic interaction between GDF9 and BMP15 is not observed in mice, consistent with the low BMP15 gene and protein expression levels in oocytes for most of folliculogenesis [109,117]. In 2004, McNatty et al. [114] hypothesized that the mechanism underlying GDF9 and BMP15 interactions is the formation of a GDF9:BMP15 heterodimer, later named cumulin [118]. It is now clear from further genetic and protein physical and functional studies that GDF9 and BMP15 act in a potent synergistic manner [102]. The current prevailing hypothesis is that GDF9: BMP15 synergism is attributed to the actions of the heterodimer cumulin [118]. It is noteworthy that the oocytes of polyovular mammals (mice, rats) express predominantly Gdf9 relative to Bmp15 (ratio 5:1), whereas monovular mammals express an equal ratio or predominantly BMP15 (GDF9:BMP15 ratios: 1.3:1 [sheep], 0.1:1 [deer] [117]). Furthermore, GDF9 in humans, and likely in all monovular species, is secreted in a latent form, in contrast to mouse GDF9 [119]. Although GDF9 is latent, it is nonetheless required for fertility, and the current hypothesis is that activation of GDF9 is achieved through formation of cumulin, in species where BMP15 is present [118] (Fig. 5). In conclusion, communication between the oocyte and somatic cells of the follicle (cumulus, granulosa and theca) via secretion of oocyte-secreted factors is an essential requirement for mammalian follicle selection and fertility. However, there is a marked species divergence in the role of GDF9 and BMP15 in these processes, which is highly unusual for TGFβ superfamily growth factors. The species-specific ovulation quota in mammals is determined by the relative bioavailability of GDF9 and BMP15 (Fig. 5). Polyovulation and high fecundity are determined

largely by a predominance of GDF9 signaling by the oocyte. In contrast, monovulation and low fecundity, such as in humans, are caused by production of active BMP15 by the oocyte, acting in concert with oocytesecreted GDF9, possibly in the form of cumulin. Together these oocyte paracrine signals modulate GC sensitivity to the gonadotropins, thereby controlling follicle selection, ovulation rate, and fecundity [114].

Oocyte Paracrine Signals Regulate GC Lineage Differentiation As follicles grow and undergo the transition from the preantral to antral stage, the GC separate into two distinct lineages: the mural granulosa cells (MGCs), which line the wall of the follicle, and the CCs (Fig. 1), which maintain an intimate association with the oocyte for the remainder of folliculogenesis. These two cell lineages perform starkly different functions from this point onward: MGCs progressively play an endocrine role, producing the major female steroids and the inhibins thereby regulating hypothalamic-pituitary function, whereas by contrast, the CCs have minimal hormone output and instead are directed to serve the needs of the developing oocyte. The cumulus-oocyte complex (COC) that ensues is a highly complex and dynamic three-dimensional structure with intricate cellular structures facilitating bidirectional communication between the oocyte and its somatic cells. Correct functioning of the COC is a prerequisite for the capacity of the oocyte to support healthy embryogenesis [120]. Given the important but differing roles of MGCs and CCs, how are these major cell lineages established and

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FIG. 5 Hypothesized mechanism regulating the species-specific ovulation quota in mammals by the relative bioavailability of GDF9, BMP15, and cumulin. Part of this research was originally published in Journal of Biological Chemistry. Mottershead DG, Sugimura S, Al-Musawi SL, Li JJ, Richani D, White MA, et al. Cumulin, an oocyte-secreted heterodimer of the transforming growth factor-beta family, is a potent activator of granulosa cells and improves oocyte quality. J Biol Chem 2015;290(39):24007–20. © The American Society for Biochemistry and Molecular Biology..

maintained? Under the dominant influence of FSH, the default pathway of GC differentiation is toward the MGC phenotype. However, within the COC, the oocyte actively prevents this pathway of differentiation by secreting potent paracrine growth factors [121,122] (Fig. 6). The oocyte establishes a morphogenic gradient of growth factors, principally through the secretion of GDF9, BMP15, and selected FGFs, which determine the CC phenotype in the oocyte’s immediate microenvironment, the effects of which dissipate with distance from the oocyte [123]. The capacity of the oocyte to regulate CC differentiation is developmentally regulated (Fig. 6). Growing oocytes in preantral follicles cannot direct CC differentiation—this capacity is first acquired following antrum formation, after oocytes have completed their growth phase [121,124]. Pharmacological ablation of oocyte-secreted GDF9 or BMP15 signaling or microsurgical removal of the oocyte from the COC causes CCs to rapidly revert to an MGC phenotype

(e.g., commencing steroid secretion) [122,125]. Hence, in order to retain the CC phenotype, the oocyte must continuously send paracrine signals to CCs to abrogate the actions of FSH driving cell differentiation toward the MGC phenotype (Fig. 6).

Impact of Oocyte-Secreted Factor Signaling on Cumulus Cell and Oocyte Function During the antral phase of folliculogenesis, the principle consequence of oocyte-secreted factor (OSF) signaling is the formation and maintenance of the COC [101]. OSFs are responsible for CCs having distinctive characteristics that contrasts those of MGCs, e.g., CCs have a high proliferation index, produce little steroids, are unresponsive to LH, are resistant to apoptosis, are highly glycolytic, and are capable of production of an extensive extracellular matrix required for ovulation [101]. Maintenance of the CC phenotype and these and other key functions

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FIG. 6 FSH and oocyte paracrine signals regulate granulosa cell lineage differentiation throughout the course of folliculogenesis. Reproduced from Scaramuzzi RJ, Baird DT, Campbell BK, Driancourt MA, Dupont J. Fortune JE, et al. Regulation of folliculogenesis and the determination of ovulation rate in ruminants. Reprod Fertil Dev 2011;23(3):444–67, with permission from CSIRO Publishing.

performed by CCs are required to support oocyte growth and development, such that the oocyte can successfully undergo fertilization and support subsequent embryo development (Fig. 7). An example of such CC-oocyte symbiosis is the fact that the oocyte has a poor capacity to metabolize glucose; instead, it instructs CCs (via BMP15 and FGF8) to upregulate glycolysis in order for CCs to supply the oocyte with substrates the oocyte needs for its own oxidative metabolism [126]. Hence, via the actions of OSFs, the oocyte is in control of its own microenvironment and its own development. A further critical message oocytes must deliver in a timely manner to CCs to facilitate selection is to instruct them to be ready for ovulation. Neither the oocyte nor CCs have appreciable LH receptors so the COC does not directly detect the LH surge, but rather receives the ovulatory signal indirectly via the epidermal growth factor (EGF)-like peptides epiregulin, amphiregulin, and betacellulin produced by the MGCs [127]. Hence, in turn, CCs need to acquire EGF family signaling capability in order to respond to the ovulatory signal. COCs in small antral follicles are unresponsive to EGF peptides and

progressively acquire responsiveness as they grow to the large antral and preovulatory stages [128]. This key developmental check point for the follicle and the oocyte is regulated by the coordinated actions of FSH with oocyte-secreted GDF9/BMP15 [129] (Fig. 8). Oocytes only acquire the capacity to enable CC EGF receptor responsiveness late in folliculogenesis [130]. This appears to be an important selection step, whereby EGF responsiveness, and thereby ovulatory functionality, is only acquired in the COC(s) of the preovulatory follicle(s), to prevent ovulation of developing oocytes in growing antral follicles that are not capable of successful fertilization or embryo development. Hence, FSH/OSF-induced EGF receptor responsiveness is a major landmark in CC differentiation, analogous to FSH-induction of LH receptors in MGCs [130,131]. Given the importance of the oocyte-somatic cell communication axis for CC function, and in turn the central role CCs play in oocyte development, it follows that OSFs impact oocyte developmental competence. Partial genetic or pharmacological ablation of oocyte-secreted GDF9 and/or BMP15 perturbs the capacity of the oocyte to

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FIG. 7 Oocyte-secreted factors are responsible for CC lineage differentiation and direct key CC functions required for oocyte developmental competence and ovulation. Reproduced from Gilchrist RB. Recent insights into oocyte-follicle cell interactions provide opportunities for the development of new approaches to in vitro maturation. Reprod Fertil Dev 2011;23 (1):23–31, with permission from CSIRO Publishing.

FIG. 8 Endocrine and oocyte paracrine signals cooperate to regulate acquisition of EGF receptor functionality and oocyte developmental competence. Reproduced from Richani D, Gilchrist RB. The epidermal growth factor network: role of in oocyte growth maturation and developmental competence. Hum Reprod Update 2018;24(1):1–14, by permission Oxford University Press; from research originally published in Sugimura S, Ritter LJ, Rose RD, Thompson JG, Smitz J, Mottershead DG, et al. Promotion of EGF receptor signaling improves the quality of low developmental competence oocytes. Dev Biol 2015;403 (2):139–49.

support development to the blastocyst stage [132,133]. Conversely, addition of exogenous OSFs to oocytes in vitro notably improves subsequent preimplantation and fetal development [132,134]. Interestingly it is only the pro-forms, and not the mature forms, of GDF9 and BMP15 that enhance oocyte quality [134,135], and furthermore, pro-cumulin is particularly potent [118]. Taken together, oocyte paracrine regulation of CC differentiation and function is vitally important for the development of the COC and for the acquisition of oocyte developmental competence (Figs. 7 and 8).

Oocyte-Somatic Cell Bidirectional Communication Maintains Oocyte Meiotic Arrest Oocytes become competent to resume meiosis between the preantral to antral stage of folliculogenesis

[136]. Spontaneous meiotic resumption occurs following removal of the COC from the follicle in most mammalian species [137]. Despite description of spontaneous meiotic resumption occurring more than 80 years ago [138], discovery of the meiotic inhibitory substance(s) derived from GCs remained elusive until 2010, and of particular interest here, was found to be coregulated by the oocyte [139]. Maintenance of meiotic arrest requires sufficient levels of intraoocyte cyclic adenosine monophosphate (cAMP) [140] (Fig. 9). However, synthesis of cAMP alone is inadequate without inhibiting its hydrolysis by the phosphodiesterase, PDE3. Inhibition of the oocyte PDE3 occurs via transfer of cyclic guanosine monophosphate (cGMP) from the somatic cells to the oocyte [140,141]. The source of cGMP and elucidation of the follicle derived “oocyte maturation inhibitor” came with the discovery of the natriuretic peptide signaling pathway in the follicle

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REFERENCES

FIG. 9 Role of OSFs in the maintenance of meiotic arrest via regulation of NPR2, IMPDH, and E2 (blue arrows). Refer to the text for description. ADCY, adenylyl cyclase; FSH, follicle-stimulating hormone; FSHR, follicle-stimulating hormone receptor; GPR3/12, G protein-coupled receptors 3 and 12; Gs, GNAS (guanine nucleotide-binding protein, α stimulating) complex.

[139]. Natriuretic peptide precursor type C (NPPC) is synthesized by GC and secreted extracellularly to activate CC natriuretic peptide receptor 2 (NPR2), a guanylyl cyclase leading to increased cGMP [139,142] (Fig. 9). Interestingly, the oocyte itself was found to play a role in controlling meiotic progression via its regulation of Npr2. Microsurgical removal of the oocyte from COCs led to a significant decrease in Npr2, while the presence of the oocyte or OSFs (specifically a combination of BMP15 and GDF9 or BMP15 and FGF8 or all three) resulted in Npr2 levels equivalent to intact COCs [139]. OSF-mediated effects on Npr2 expression also require estradiol [143]. While OSFs have no effect on Nppc expression, estradiol does [144], which interestingly is regulated by OSFs [145]. The influence of the oocyte on the NPPC/ NPR2 system and meiotic arrest is further strengthened by the finding that the rate-limiting enzyme for GTP synthesis (precursor for cGMP), inosine monophosphate dehydrogenase, is also regulated by OSFs [146] (Fig. 9). Thus, in a modification to the original hypothesis of granulosa controlled meiotic arrest, discoveries in recent years have shed light on the requirement of the oocyte in regulating the NPPC/NPR signaling system in GC and CC, which in turn maintain oocyte meiotic arrest prior to the LH-surge [146].

CONCLUSIONS Follicle selection occurs at all stages of folliculogenesis from the primordial follicle resting in the ovarian reserve to the dominant ovulatory follicle, and it involves both the oocyte and the CC, GC, and theca somatic cells of

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the follicle. Selection can be thought of as a regulated outcome between survival and growth or death, predominately through atresia, which determines which follicles containing an oocyte survive and eventually ovulate. Selection depends on a follicle being at the appropriate place and stage of development to be able to respond to the milieu within the niche. In particular, selection depends on the check points controlled by the oocyte. In the primordial follicle where the death process originates in the oocyte, it is a balance between DNA repair and death. The death process in primordial oocytes includes atresia but may also involve other forms of demise such as autophagy. In growing follicles, the atresia originates in the follicular cells that can no longer support folliculogenesis with subsequent demise of the follicle, including the oocyte. At all stages of folliculogenesis, the oocyte plays a significant role in selection. There are still many unanswered questions about folliculogenesis that provide opportunities for future research. For example, • Why is the ovarian reserve so large when only a small percentage of the follicles ever ovulate, even in polyovular species? • What governs the balance between DNA repair and cell death in the oocyte? • What determines the rate of selection of primordial follicles for activation and growth? • Do mitochondria play a role in follicle selection? • What are the mechanisms by which partial actions of GDF9/BMP15 result in an increase in ovulation rate, but inaction of these factors induces sterility? • Are there other growth factors that are essential for survival and selection of follicles?

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FURTHER READING

[117] Crawford JL, McNatty KP. The ratio of growth differentiation factor 9: bone morphogenetic protein 15 mRNA expression is tightly co-regulated and differs between species over a wide range of ovulation rates. Mol Cell Endocrinol 2012;348(1):339–43. [118] Mottershead DG, Sugimura S, Al-Musawi SL, Li JJ, Richani D, White MA, et al. Cumulin, an oocyte-secreted heterodimer of the transforming growth factor-beta family, is a potent activator of granulosa cells and improves oocyte quality. J Biol Chem 2015;290(39):24007–20. [119] Simpson CM, Stanton PG, Walton KL, Chan KL, Ritter LJ, Gilchrist RB, et al. Activation of latent human GDF9 by a single residue change (Gly 391 Arg) in the mature domain. Endocrinology 2012;153(3):1301–10. [120] Gilchrist RB, Ritter LJ, Armstrong DT. Oocyte-somatic cell interactions during follicle development in mammals. Anim Reprod Sci 2004;82-83:431–46. [121] Eppig JJ, Wigglesworth K, Pendola F, Hirao Y. Murine oocytes suppress expression of luteinizing hormone receptor messenger ribonucleic acid by granulosa cells. Biol Reprod 1997;56(4):976–84. [122] Li R, Norman RJ, Armstrong DT, Gilchrist RB. Oocyte-secreted factor(s) determine functional differences between bovine mural granulosa cells and cumulus cells. Biol Reprod 2000;63(3):839–45. [123] Hussein TS, Froiland DA, Amato F, Thompson JG, Gilchrist RB. Oocytes prevent cumulus cell apoptosis by maintaining a morphogenic paracrine gradient of bone morphogenetic proteins. J Cell Sci 2005;118(Pt 22):5257–68. [124] Gilchrist RB, Ritter LJ, Armstrong DT. Mouse oocyte mitogenic activity is developmentally coordinated throughout folliculogenesis and meiotic maturation. Dev Biol 2001;240(1):289–98. [125] Dragovic RA, Ritter LJ, Schulz SJ, Amato F, Thompson JG, Armstrong DT, et al. Oocyte-secreted factor activation of SMAD 2/3 signaling enables initiation of mouse cumulus cell expansion. Biol Reprod 2007;76(5):848–57. [126] Sugiura K, Su YQ, Diaz FJ, Pangas SA, Sharma S, Wigglesworth K, et al. Oocyte-derived BMP15 and FGFs cooperate to promote glycolysis in cumulus cells. Development 2007;134(14):2593–603. [127] Park JY, Su YQ, Ariga M, Law E, Jin SL, Conti M. EGF-like growth factors as mediators of LH action in the ovulatory follicle. Science 2004;303(5658):682–4. [128] Prochazka R, Srsen V, Nagyova E, Miyano T, Flechon JE. Developmental regulation of effect of epidermal growth factor on porcine oocyte-cumulus cell complexes: nuclear maturation, expansion, and F-actin remodeling. Mol Reprod Dev 2000; 56(1):63–73. [129] Diaz FJ, Wigglesworth K, Eppig JJ. Oocytes are required for the preantral granulosa cell to cumulus cell transition in mice. Dev Biol 2007;305(1):300–11. [130] Ritter LJ, Sugimura S, Gilchrist RB. Oocyte induction of EGF responsiveness in somatic cells is associated with the acquisition of porcine oocyte developmental competence. Endocrinology 2015;156(6):2299–312. [131] Sugimura S, Ritter LJ, Rose RD, Thompson JG, Smitz J, Mottershead DG, et al. Promotion of EGF receptor signaling improves the quality of low developmental competence oocytes. Dev Biol 2015;403(2):139–49. [132] Hussein TS, Thompson JG, Gilchrist RB. Oocyte-secreted factors enhance oocyte developmental competence. Dev Biol 2006;296 (2):514–21. [133] Su YQ, Wu X, O’Brien MJ, Pendola FL, Denegre JN, Matzuk MM, et al. Synergistic roles of BMP15 and GDF9 in the development and function of the oocyte-cumulus cell complex in

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mice: genetic evidence for an oocyte-granulosa cell regulatory loop. Dev Biol 2004;276(1):64–73. Sudiman J, Ritter LJ, Feil DK, Wang X, Chan K, Mottershead DG, et al. Effects of differing oocyte-secreted factors during mouse in vitro maturation on subsequent embryo and fetal development. J Assist Reprod Genet 2014;31(3):295–306. Sudiman J, Sutton-McDowall ML, Ritter LJ, White MA, Mottershead DG, Thompson JG, et al. Bone morphogenetic protein 15 in the pro-mature complex form enhances bovine oocyte developmental competence. PLoS One 2014;9(7):e103563. Sorensen RA, Wassarman PM. Relationship between growth and meiotic maturation of the mouse oocyte. Dev Biol 1976;50(2):531–6. Edwards RG. Maturation in vitro of mouse, sheep, cow, pig, rhesus monkey and human ovarian oocytes. Nature 1965; 208(5008):349–51. Pincus G, Enzmann EV. The comparative behavior of mammalian eggs in vivo and in vitro: I. The activation of ovarian eggs. J Exp Med 1935;62(5):665–75. Zhang M, Su YQ, Sugiura K, Xia G, ligand EJJGc. NPPC and its receptor NPR2 maintain meiotic arrest in mouse oocytes. Science 2010;330(6002):366–9. Vaccari S, Weeks 2nd. JL, Hsieh M, Menniti FS, Conti M. Cyclic GMP signaling is involved in the luteinizing hormone-dependent meiotic maturation of mouse oocytes. Biol Reprod 2009; 81(3):595–604. Norris RP, Ratzan WJ, Freudzon M, Mehlmann LM, Krall J, Movsesian MA, et al. Cyclic GMP from the surrounding somatic cells regulates cyclic AMP and meiosis in the mouse oocyte. Development 2009;136(11):1869–78. Geister KA, Brinkmeier ML, Hsieh M, Faust SM, Karolyi IJ, Perosky JE, et al. A novel loss-of-function mutation in Npr2 clarifies primary role in female reproduction and reveals a potential therapy for acromesomelic dysplasia, Maroteaux type. Hum Mol Genet 2013;22(2):345–57. Zhang M, Su YQ, Sugiura K, Wigglesworth K, Xia G, Eppig JJ. Estradiol promotes and maintains cumulus cell expression of natriuretic peptide receptor 2 (NPR2) and meiotic arrest in mouse oocytes in vitro. Endocrinology 2011;152(11):4377–85. Lee KB, Zhang M, Sugiura K, Wigglesworth K, Uliasz T, Jaffe LA, et al. Hormonal coordination of natriuretic peptide type C and natriuretic peptide receptor 3 expression in mouse granulosa cells. Biol Reprod 2013;88(2):42. Vanderhyden BC, Macdonald EA. Mouse oocytes regulate granulosa cell steroidogenesis throughout follicular development. Biol Reprod 1998;59(6):1296–301. Wigglesworth K, Lee KB, O’Brien MJ, Peng J, Matzuk MM, Eppig JJ. Bidirectional communication between oocytes and ovarian follicular somatic cells is required for meiotic arrest of mammalian oocytes. Proc Natl Acad Sci USA 2013;110(39): E3723–9.

Further Reading [147] Gilchrist RB. Recent insights into oocyte-follicle cell interactions provide opportunities for the development of new approaches to in vitro maturation. Reprod Fertil Dev 2011;23(1):23–31. [148] Richani D, Gilchrist RB. The epidermal growth factor network: role of in oocyte growth maturation and developmental competence. Hum Reprod Update 2018;24(1):1–14.

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C H A P T E R

2 Regulation of Follicle Formation and Development by Ovarian Signaling Pathways Rexxi D. Prasasya Kelly E. Mayo Introduction

demonstrate the diversity of signaling pathways used in intrafollicular cell communication. During embryonic development, primordial germ cells (PGCs) arise at the proximal epiblast and proceed to proliferate, migrate, and eventually colonize the bipotential gonad [1]. While mechanisms of sex determination are beyond the scope of this chapter, it is now known that the process is somatic cell driven, and ovarian fate is promoted in the absence of the Y chromosome and Sry gene expression. In the XY individual, the presence of SRY suppresses Wnt4/β-catenin pathway signaling (reviewed in Ref. [2]). Conversely, in the XX individual, the lack of SRY allows for initiation of pregranulosa cell differentiation as well as continued suppression of the Sertoli cell fate by an increase in Wnt4/β-catenin signaling (reviewed in Ref. [2]). At around embryonic day 13.5 (E13.5) in the mouse or 10 weeks of gestation in the human, a rise in retinoic acid (RA) levels within the embryonic ovary induces the expression of stimulated by retinoic acid gene 8 (Stra8), which triggers germ cells, or oocytes, to enter meiosis [3,4]. RA in the fetal gonad originates from the mesonephros, and is actively degraded by the P450 cytochrome enzyme Cyp26b1 in the embryonic testis, thus preventing early entry into meiosis in spermatogonia [5]. In contrast, Cyp26b1 expression is downregulated by E12.5 in fetal mouse ovaries, thus allowing the rise in RA and initiation of meiosis [4,5]. Interestingly, meiosis, while necessary for fertility, does not appear to be required for oocyte differentiation or the ability of oocytes to interact with granulosa cells to form follicles. While there is an increased rate of germ cells loss in Stra8-deficient ovaries, leading to complete

In mammals, ovarian follicles make up the basic functional unit of the female gonad, the ovary. Within the follicle, the female germ cell, the oocyte, interacts with the surrounding granulosa cells, and they in turn with thecal cells, through both physical contact and secreted signals. Bidirectional communication between the oocyte and the follicular somatic cells is necessary to ensure fertility through the eventual ovulation of a fertilizable oocyte as well as production of hormones by the somatic cells that are important for overall physiology. The endocrine control of follicular growth and ovulation by the pituitary gonadotropins follicle-stimulating hormone (FSH) and luteinizing hormone (LH) is well recognized from classic physiology experiments. More recently, the advent of modern molecular biology techniques and genetic mouse models has allowed investigations into the cellular signaling pathways and gene regulatory events that are responsible for follicle formation and development. As will become clear early in this chapter, a myriad of overlapping and often interacting pathways govern each specific stage of follicular development. It is perhaps by design that a built-in redundancy exists in the molecular control of a physiological process that ultimately determines the continuation of the species. We begin by focusing our discussion on emerging signaling pathways that contribute to the least understood stages of follicular development, the formation and activation of primordial follicles. Subsequently, we review gonadotropinindependent and -dependent follicular growth by focusing on several types of dominant local factors that

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germ cell depletion by 6–8 weeks of age, the surviving germ cells are contained within follicular structures, which can be ovulated through exogenous hormone stimulation [6]. Moreover, these 2n oocyte-like cells are able to produce a zona pellucida (ZP) matrix as would bona fide oocytes [6]. Meiosis will continue to the diplotene stage of meiotic prophase I, at which it arrests and will not be resumed until postovulation. Following meiotic arrest, germ cell clusters, which arise through incomplete cytokinesis of rapidly dividing germ cells, begin interacting with surrounding pregranulosa cells to form a structure termed a germ cell syncytia, germ cell cyst, or germ cell nest, which serves as a precursor to primordial follicles [7]. Pregranulosa cells are thought to arise from at least three distinct origins: somatic cells within the bipotential gonad, the ovarian surface epithelium, and a structure at the ovarymesonephros border termed the rete ovarii [2,8,9]. The number of primordial follicles formed following completion of germ cell nest breakdown contributes to establishing the reproductive life span of the individual. The dynamics of primordial follicle formation and activation were recently studied by two independent groups using highly sensitive and temporally inducible reporter lines to label granulosa cells [10,11]. These studies show that while some primordial follicles are recruited into the growing population immediately following their formation, the majority of primordial follicles remain quiescent for varying periods of time [10,11]. Mork and colleagues confirmed that in the mouse, there are at least

two different sources of pregranulosa cells, one migrating from the mesonephros and one migrating from the surface epithelium [11]. Moreover, migration and the subsequent differentiation and association with germ cell nests by these two distinct types of pregranulosa cells occur sequentially, resulting in what is now accepted to be two waves of primordial follicle formation [10,11]. Both studies demonstrate that following birth, primordial follicles with embryonically labeled granulosa cells (presumably part of the first wave of formation) are immediately recruited into the growing follicle pool, some reaching the early antral stage by PND23 [10,11]. Furthermore, these first wave follicles appear to contribute to fertility in early adulthood, as labeled corpora lutea (CLs) are detected in PND90 ovaries [10]. These studies demonstrate that early events such as follicular formation and activation have direct consequences on fertility during adulthood.

CELLULAR SIGNALS REGULATING GERM CELL NEST BREAKDOWN AND PRIMORDIAL FOLLICLE FORMATION A mammalian female is born with a finite number of oocytes, and as such, the subsequent number of primordial follicles formed will in part determine the fecundity and reproductive life span of the individual. Fig. 1 illustrates the process of follicle formation and development,

FIG. 1 Formation and development of the ovarian follicle. During gestation, migrating primordial germ cells form connected clusters through incomplete cytokinesis and aggregation. Following colonization of the embryonic ovary, somatic pregranulosa cells will interact with these germ cell clusters to form germ cell nests. Primordial follicles are formed when pregranulosa cells invade the germ cell clusters to encapsulate single oocytes within a follicle. This process is accompanied by programmed oocyte death, leaving only about 30% surviving oocytes at the completion of nest breakdown. Promoters of nest breakdown include paracrine and juxtacrine signaling through Activin, KITL, NGF, and Notch pathway signaling. Conversely, nest breakdown is inhibited by steroid hormones such as estrogen and progesterone. Primordial follicles are gradually recruited into the growing pool throughout the reproductive lifetime. The extracellular cues that initiate primordial follicle activation remain unclear; however, PI3K/AKT and mTORC signaling within the oocyte are known to govern its exit from quiescence. The growing follicle population, in turn, provides negative feedback to the primordial follicle pool through production of AMH. Oocyte activation is also regulated by a network of oocyte-specific transcription factors that includes NOBOX, LHX8, SOHLH1, and SOHLH2. These transcription factors regulate expression of oocyte-specific genes including those involved in ZP matrix deposition and Gdf9 and Bmp15, which encode secreted factors that are necessary for continued follicle growth. As the follicle reaches the multilayer secondary stage, the final somatic cell layer, the theca layer, is specified through activation of Hh signaling. Up to this stage, follicle development is independent of pituitary gonadotropins. Following formation of the fluid-filled antral cavity, FSH becomes dominant in regulating proliferation and differentiation of granulosa cells toward the preovulatory phenotype working in concert with local paracrine signals.

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accompanied by major factors that are involved in each developmental stage. In the XX individual, PGCs undergo rapid mitotic division as they migrate and colonize the gonad until the initiation of meiosis [12]. This mitotic division is accompanied by incomplete cytokinesis [13–15], resulting in structures termed nests or syncytia where multiple cells are connected by cytoplasmic bridges, similar to those found in invertebrates [16,17]. There is evidence that organelles, including centrosomes, Golgi bodies, and mitochondria, are transferred between sister oocytes through these cytoplasmic bridges, producing a cloud of organelles termed the Balbiani body within oocytes that are more likely to survive following cyst resolution [18,19]. While there is a strong correlation between the presence of the Balbiani body and oocyte survival [18], the cytoplasmic bridges do not appear to be absolutely necessary for female fertility. This is demonstrated by a knockout model of Tex14, a component of the cytoplasmic bridges [20]. While Tex14
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mice are endowed with reduced numbers of oocytes as compared to littermate controls following completion of germ cell nest breakdown, their primordial follicle pool is able to support a normal reproductive life span and full fertility [20]. Primordial follicles are formed when pregranulosa cells send projections toward the germ cell nests to encapsulate an individual oocyte within a follicle [7]. In the mouse, germ cell nest breakdown begins shortly before birth and concludes at about PND6, when more than 80% of germ cells are contained within primordial follicles [7,21]. Only about 30% of oocytes from germ cell nests survive to become primordial follicles, as a wave of oocyte death occurs concurrently with the progression of germ cell nest breakdown [7]. It is hypothesized that this massive loss of oocytes allows for the optimization of the number of available pregranulosa cells to the number of surviving oocytes, and also for the remaining oocytes to acquire organelles from those that are lost. Alternatively, oocyte death has also been suggested to directly facilitate the resolution of germ cell syncytia into primordial follicles, by creating smaller syncytia following selective oocyte lost [7], or as a means to eliminate oocytes with chromosomal aberrations [22]. There is no consensus on whether germ cell nest breakdown is driven by the germ cells or by the somatic cells. It is apparent that interactions between properly specified somatic pregranulosa cells and oocytes are important for successful nest breakdown and establishment of the primordial follicle pool. Foxl2, a gene encoding a winged-helix forkhead gene transcription factor, is one of the earliest markers identified to be required for granulosa cell specification [23]. Foxl2
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mice are infertile due to incompetency of pregranulosa cells to interact with germ cells clusters to form follicles [24]. In turn, developmental competency of the oocyte is also important in

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primordial follicle formation. For example, an ovary that lacks expression of the oocyte-restricted Factor in germ line alpha (Figla), a basic helix-loop-helix transcription factor that is required for ZP formation [25], does not form follicles, leading to eventual demise of oocytes within disorganized germ cell nests [24]. In the following sections, we discuss the proposed mechanisms for programmed germ cell death during the period of nest breakdown, as well as emerging signaling pathways that contribute to the formation of primordial follicles.

Molecular Control of Neonatal Oocyte Survival An experimental tool that has been extensively used to study signaling pathways during germ cell nest breakdown is the ex vivo culture of neonatal rodent ovaries [26]. This organotypic culture system, along with genetic mouse models, has revealed many candidate factors and signaling pathways that contribute to bidirectional communication between somatic pregranulosa cells and oocytes during nest breakdown. Several investigations have explored whether modulation of oocyte survival can alter the primordial follicle endowment, and thus the reproductive life span. Of interest is the B cell lymphoma-2 (BCL-2) family of proteins, whose members are key regulators of cellular apoptosis. These proteins promote or inhibit apoptosis by controlling mitochondrial outer membrane permeabilization. Deletion of Bax, a proapoptotic member of the Bcl-2 family, prolongs reproductive life span by 10–12 months in the mouse due to a decreased rate of follicular atresia [27]. While it was initially thought that the primordial follicle pool is not altered in Bax
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ovaries, a second study shows an increase in the absolute number of germ cells, and of primordial follicles formed, in embryonic and neonatal Bax
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ovaries [28]. Surprisingly, a wave of oocyte death during the period of germ cell nest breakdown is still observed in Bax
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ovaries, suggesting that oocyte death during nest breakdown is Bax-independent. Additional evidence that suggests alternative mechanisms of apoptotic pathway activation in lieu of BCL-2 family proteins in neonatal oocytes comes from knockouts of Puma and Bcl-2-modifying factor (Bmf), both proapoptotic proteins [29,30]. Similar to the Bax
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ovary, both Puma
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and Bmf
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ovaries contain more oocytes at embryonic and neonatal ages [29,30]. In both models, however, a wave of germ cell death occurs during the period of nest breakdown. In the case of Bmf
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ovaries, the greater number of germ cells does not result in a larger endowment of primordial follicles at the conclusion of germ cell nest breakdown [30]. This normalization in the number of follicles is also observed in an overexpression model of the antiapoptotic gene Bcl2 where the increased numbers of primordial follicles formed

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are eventually lost, with normalization to that of littermate controls by early adulthood [31]. While the mechanisms behind the initiation of programmed oocyte death during nest breakdown remain unresolved, it is generally accepted that oocyte death occurs via apoptosis following activation of CASPASE-2. Activation of caspases or cysteine-aspartic proteases leads to cleavage of cellular proteins and triggers apoptosis. At PND4, Caspase2
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ovaries contain greater number of primordial follicles, and thus oocytes, compared to ovaries of littermate controls [32]. Moreover, Caspase2
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oocytes are shown to better survive assault from exposure to the chemotherapeutic agent doxorubicin [32]. While genetic mouse models have allowed for identification of necessary as well as dispensable molecules in programmed oocyte death, the mechanisms by which surviving oocytes are able to escape apoptosis remain elusive.

The Role of Steroid Hormones in Nest Breakdown In rodents, cattle, and macaques, the period of nest breakdown coincides with a drastic drop in the levels

of steroid hormones to which the newborns or fetuses are exposed [33–37], leading to the hypothesis that estrogen and progesterone act as negative regulators of primordial follicle assembly (Fig. 2). Using ex vivo cultures of neonatal rat and mouse ovaries, estradiol and progesterone were found to inhibit primordial follicle formation, resulting in the retention of germ cell nests [38,39]. The effect of progesterone on nest breakdown does not appear to be due to its conversion to estradiol, since exposure of neonatal ovaries to a nonhydrolyzable form of progesterone, promegestone, also results in retention of nests [39]. Estrogen receptor-α (Esr1) is expressed in pregranulosa cells, and estrogen receptor-β (Esr2) is expressed in both pregranulosa cells and oocytes at birth in mouse ovaries [40,41]. Using selective agonists and antagonists for ER-α, ER-β, and the nongenomic, membrane-bound form of estrogen receptor, Chen and colleague show that inhibition of germ cell nest breakdown by estrogen can be carried out by all three forms of estrogen receptor [40]. It is thought that there is a critical window within which nest breakdown must occur, after which pregranulosa cell invasion ceases regardless of whether all oocytes have been contained in a follicle. Therefore, it is

FIG. 2

Select signals that promote germ cell nest breakdown and primordial follicle formation. The process of germ cell nest breakdown occurs during midgestation in human and during the neonatal period in the mouse. Experimental evidence has shown that high levels of estrogen and progesterone inhibit primordial follicle formation. Several signaling pathways that are initiated by growth factors have been shown to promote primordial follicle formation. Activin stimulates proliferation of pregranulosa cells and promotes Notch signaling in the granulosa cells, which enhances primordial follicle formation. The Ntrk receptors are localized to both oocytes and pregranulosa cells during the period of nest breakdown and their ligands are expressed by the pregranulosa cells. Signaling through Ntrks promotes nest breakdown, potentially through upregulation of Jag1 expression in the oocyte. The juxtacrine signal Notch, which is mediated by the ligand JAG1 in the oocyte and the receptor NOTCH2 in pregranulosa cells, is emerging as an important promoter of nest breakdown and granulosa cell proliferation in the early stages of follicle development. Signaling through the c-KIT receptor in the oocyte and its ligand KITL in the granulosa cells promotes follicle assembly through its contribution to programmed germ cell death.

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postulated that remnants of germ cell nests from delayed breakdown will ultimately lead to the formation of multioocytic follicles (MOFs). The in vitro findings of the negative effects of estrogen on nest breakdown are consistent with in vivo studies where neonatal exposure to the synthetic estrogen diethylstilbestrol (DES) [42,43] or the soy phytoestrogen genistein [44,45] results in delayed germ cell nest break down and a higher incidence of MOFs as compared to control mice. The consequence of MOFs for fertility remains unclear. Iguchi et al. report reduced fertilization capacity of oocytes obtained from ovulated MOFs in a mouse model, but the confounding effects of early exposure to DES in these oocytes cannot be eliminated [46]. However, at least one report suggests that using oocytes from MOFs does not lead to any adverse outcomes in human in vitro fertilization [47].

The Roles of Growth Factors and Somatic CellCycle Progression During Germ Cell Nest Breakdown Several growth factors have been implicated in germ cell nest breakdown. Activin is a member of the TGF-β family of proteins that is expressed in granulosa cells of growing follicles [48]. Treatment of neonatal mice with activin leads to a 30% increase in the number of primordial follicles, and this is associated with increased proliferation of both granulosa cells, and, to a limited degree, the germ cells population (Fig. 2) [21]. Similar to several genetic models of BCL-2 family protein modulation described earlier, the expanded primordial follicle pool that is observed in neonatally activin-exposed ovaries is not maintained into reproductive maturity, a paradigm that is described as an inherent quorum-sensing mechanism by the ovary [21]. Follistatin (FST) is expressed in the gonad and functions to bind activin and neutralize its action [49]. Germ cell nest breakdown is significantly delayed in a mouse model that selectively expresses a high-affinity isoform of FST (FST-288) due to an increase in somatic cell proliferation and a decrease in germ cell death during the neonatal period [50]. Whether this delay in germ cell nest breakdown leads to MOFs was not characterized [51]; however, the primordial follicle pool is expanded in the FST-288 mouse [50]. Despite this, the FST-288 mouse is subfertile due to accelerated depletion of the primordial follicle pool in adulthood [52]. Consistent with this genetic mouse model, the addition of FST288 to ex vivo ovarian culture system during the period of germ cell nest breakdown leads to a delay in primordial follicle formation [53]. FST-288 treatment leads to reduced granulosa cell proliferation and suppressed Notch signaling, a pathway that is known to promote germ cell nest breakdown, in granulosa cells [53]. These studies illustrate the importance of activin signaling in

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formation and maintenance of the primordial follicle pool through its influence on granulosa cell proliferation and likely, germ cell survival. The importance of receptor tyrosine kinase (RTK) signaling in promoting germ cell nest breakdown is beginning to be elucidated. Kit ligand (KITL) and its tyrosine-kinase receptor c-KIT are localized to pregranulosa cells and oocytes, respectively, during ovarian embryonic development (Fig. 2) and have been shown to be necessary for PGC migration [54]. Their expression remains during the period of nest breakdown and exposure of cultured neonatal mouse ovaries to a c-KIT blocking antibody leads to the retention of germ cell nests, which is hypothesized to result from inhibition of oocyte death [55]. Furthermore, exposure of neonatal rats to the small molecule tyrosine kinase inhibitor imatinib, which targets several RTKs including c-KIT, leads to retention of germ cell nests and an increased incident of MOFs [56]. Expression of neurotrophic tropomyosin-related kinase (Ntrk) receptors, a family of RTKs, and their soluble neurotrophin ligands is dynamically regulated during the period of germ cell nest breakdown in neonatal rat ovaries [57]. Knockout mouse models of the ligand nerve growth factor (Ngf) or the receptors Ntrk1 and Ntrk2 lead to a delay in germ cell nest breakdown and reduction in the number of primordial follicles formed [58,59]. Further evidence for the involvement of Ntrk family of receptors in nest breakdown is demonstrated by the enhancement of primordial follicle formation observed in neonatal mouse ovaries that are cultured with connective tissue growth factor (CTGF), whose expression is upregulated during the neonatal period [60]. While the receptor by which CTGF exerts its effects during germ cell nests breakdown has not been specified, there is evidence of its ability to activate NTRK1 in human mesenchymal cell [61]. Additional evidence for an association between regulations of germ cell nest breakdown and cell cycle progression comes from a knockout model of the cell-cycle inhibitor p27KipI. A member of the Cip/Kip subfamily, p27KipI, acts as a cyclin-dependent kinase inhibitor. p27KipI is localized to somatic cells of primordial follicles, and p27KipI
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ovaries show an acceleration in germ cell nest breakdown, reaching completion by PND2 [62]. While proliferation was not directly measured, this result suggests that the loss of inhibition of somatic cell proliferation facilitates the formation of primordial follicles. This is consistent with the correlation between granulosa cell proliferation and germ cell nest breakdown that is observed following modulation of activin signaling. The number of primordial follicles formed in p27KipI
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ovaries is double that of litter-mate controls; however, the number of follicles eventually normalizes by early adulthood following massive follicular attrition, consistent with the quorum-sensing model proposed to govern

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follicular dynamic in the ovary [63]. p27KipI also appears to control primordial follicle maintenance, which we will discuss in more detail in the subsequent section, as p27KipI
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mice are infertile due to a premature depletion of the primordial follicle pool [62].

Regulation of Germ Cell Nest Breakdown by Notch Signaling To form primordial follicles, pregranulosa cells must invade germ cells clusters and form physical contacts with the oocyte that will eventually be surrounded and survive. While direct contact between the first layer of granulosa cells and the oocyte is initially unimpeded, ZP matrix deposition following primordial follicle activation will eventually limit these continuous cell-to-cell contacts. To maintain physical contact with the oocyte, granulosa cells are able to send cytonemes that traverse the ZP and form both adherens and gap junctions with the oocyte plasma membrane, termed transzonal projections (TZPs) [64]. While initially thought to exclusively function as a means for granulosa cells to shuttle nutrients in response to the metabolic demands of the oocyte during development, the presence of TZPs provides a rationale for investigations into the potential role of cell-contact-dependent signaling during folliculogenesis. Among these, the highly conserved Notch juxtacrine signaling pathway has emerged as a key regulator of germ cell nest breakdown. In mammals, Notch signaling occurs when one of the five-membrane-bound ligands (JAGGED1-2, DLL1,3,4) bind to one of the fourmembrane-bound receptors (NOTCH1-4) on an opposing cell [65]. Ligand binding leads to conformational changes of the Notch receptor, initiating a proteolytic cascade that will ultimately lead to a γ-secretase-dependent cleavage and liberation of the Notch receptor intercellular domain (NICD) [66]. NICD will translocate into the nucleus and function as a transcriptional coactivator of Notch target genes upon binding with the obligate cofactor RBPJ [67]. Notch signaling is known to regulate many cell-fate decisions during development [68], and to control the proliferation and differentiation of somatic follicle cells (analogous to somatic pregranulosa cells in mammals) during ovariole development in the Drosophila melanogaster ovary [69]. Notch receptors and ligands are expressed in neonatal and adult ovaries of mice [70–72]. Using a transgenic Notch reporter mouse line in which the green fluorescent protein (Gfp) is expressed in an NICD/RBPJ-dependent manner [73], Notch active cells are detected as early as E15.5 in the mouse ovary [74]. By E18.5, Notch activity is detected in granulosa cells that are actively reorganizing to encapsulate germ cells clusters [74]. Reporter activity and Notch receptor expression continue in somatic

granulosa cells as follicles are activated and mature. The oocyte expresses predominantly the ligand JAG1 at this time. The importance of functional Notch signaling was demonstrated using an ex vivo ovarian culture system. Following a culture period that represents the period of germ cell nest breakdown, treatment of neonatal mouse ovaries with the γ-secretase inhibitor, DAPT, leads to the retention of germ cell nests and formation of fewer primordial follicles as compared to the vehicle controls [75,76]. DAPT treatment also leads to suppression of oocyte-specific transcription factors that are known to be important for early follicular development such as newborn ovary homeobox protein (Nobox), Figla, spermatogenesis and oogenesis specific basic helix-loop-helix 2 (Sohlh2), and LIM/homeobox protein 8 (Lhx8) [77]. A disintegrin and metalloproteinase 10 (ADAM-10) is an integral component of the proteolytic cleavage sequence that leads to the liberation of NICD [78]. In vitro treatment of embryonic ovaries with an ADAM10 inhibitor or conditional deletion of ADAM-10 in somatic pregranulosa cells leads to suppression of Notch signaling (as identified by reduced level of NICD and decreased expression of Notch target genes Hey2 and Hes1) and retention of germ cell nests [79]. In other mouse models, conditional deletion, using the Cre-loxP system, of the most abundantly expressed Notch receptor, Notch2, in the pregranulosa cells (N2KO) leads to decreased numbers of primordial follicles and formation of MOFs [74,80]. The rate of apoptosing oocytes is significantly decreased at PND1 in N2KO ovaries, which is associated with delayed germ cells nests breakdown [80]. A null mutation in the Notch receptor modifier Lunatic fringe (Lfng) also reveals the presence of MOFs and infertility [81]. Overall, these studies demonstrate the importance of canonical Notch signaling in the somatic pregranulosa cells for proper follicular formation. As expected, conditional deletion of the most abundantly expressed ligand Jag1, from the oocytes (J1KO) phenocopies, in a slightly more robust manner, the N2KO line [74]. Jag1 expression within the oocyte was recently described to be controlled by RAC1, a small GTPase switch that affects gene transcription via the STAT3 pathway (Fig. 2) [82]. Inhibition of RAC1 leads to decreased numbers of primordial follicles formed and reduced expression of several oocyte specific factors, including growth differentiation factor-9 (Gdf9), bone morphogenic protein-15 (Bmp15), Nobox, and Jag1 [82]. MOFs in N2KO and J1KO mice can be extremely large, containing many oocytes, suggesting that they are indeed remnants of incompletely broken down germ cells nests [74]. Both the N2KO [80] and J1KO [74] lines are subfertile, indicating that disruption of critical signals during follicular formation may lead to adverse fertility outcomes. Together, these genetic mouse lines support the

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model that physical contact between oocyte and pregranulosa cells facilitates initiation of Notch signaling, which in turn acts as a critical mean of bidirectional communication between the two cell types during follicular assembly (Fig. 2). It is interesting to note that many of the previously discussed candidates for regulators of germ cell nest breakdown appear to function upstream of Notch signaling. Progesterone exposure of embryonic and neonatal ovaries, which is shown to suppress follicular assembly, inhibits expression of Jag2, Notch2, and a known Notch target gene Hey2 [83]. Jag1, Hes1, and Hey2 were identified to be significantly downregulated in neonatal Ntrk2
/
ovaries, in which reductions in follicular assembly and granulosa cell proliferation are observed [84]. Additionally, lentiviral-mediated targeted expression of Jag1 in oocytes of Ntrk2
/
ovaries was shown to rescue the defects in granulosa cell proliferation [84]. Fig. 2 summarizes known as well as proposed interactions between different signaling pathways that regulate germ cell nest breakdown. It will require further exploration to understand whether Notch signaling acts as a potential integrator of signals that promote germ cell nest breakdown in the mammalian ovary.

REGULATION OF PRIMORDIAL FOLLICLE ACTIVATION Throughout the reproductive life span, growth is initiated in a portion of primordial follicles, and they transition to the activated primary follicle stage. Morphologically, this is characterized by the transition of the single layer of squamous granulosa cells into cuboidal cells, as well as deposition of a ZP matrix by the oocyte [85,86]. In turn, a cohort of these small activated follicles will be recruited to undergo further growth during each reproductive cycle, with their final fates being either atresia or ovulation. Perhaps one of the least understood aspects of the mammalian ovary is the physiological cues that are responsible for initiating the activation of some, but not all, primordial follicles at any given moment. One candidate for a signal regulating primordial follicle activation is the Anti-M€ ullerian hormone (AMH). A member of the TGF-β superfamily of proteins, AMH is produced by granulosa cells of growing follicles in the ovary [87,88]. Clinically, AMH is used as a marker of ovarian follicular reserve due to the well-characterized positive correlation between its serum levels and antral follicle counts [89]. Serum levels of AMH decline with age, and AMH levels are often found to be low or undetectable in clinical cases of primary ovarian insufficiency [90,91]. Female mice with a null mutation of Amh are fertile; however, more growing follicles are detected in adult

29

Amh
/
ovaries compared to wild-type littermates [92]. By 4 months of age, the number of primordial follicles is significantly reduced in Amh
/
ovaries, becoming almost completely depleted by 13 months of age [93]. Despite the increase in the absolute number of growing follicles, Amh-/- ovaries do not contain an increased proportion of preovulatory follicles due to an enhancement in the rate of atresia of small preantral follicles [94]. Conversely, addition of AMH to ex vivo cultured ovaries leads to a decrease in the growing follicle population by about 40% [93]. These studies suggest that the growing follicle population plays a role in regulating the dynamics of activation of the primordial follicle pool through production of AMH, which acts as a maintenance signal. Despite unresolved questions regarding the extracellular signals initiating activation, studies involving the intracellular pathways regulation of follicle activation have been significantly advanced, largely through the use of mouse genetic models. Fig. 3 summarizes phenotypes of relevant mouse genetic models that are discussed in the proceeding sections. In addition to discussing pathways intrinsic to the oocyte and their role in primordial follicle activation, we review below the networks of transcriptional control associated with oocyte and granulosa cell differentiation during the transition from the primordial to the primary follicle stage.

Intracellular Signaling Pathways That Regulate Primordial Follicle Activation Regulation of Oocyte Activation by the PI3K/AKT Pathway Perhaps the most intensely investigated pathway in primordial follicle activation is PI3K/AKT signaling within the oocyte. A major kinase cascade that can be activated by RTKs, the PI3K/AKT pathway (Fig. 4) functions to regulate metabolism, growth, proliferation, and survival through activation of gene transcription or protein synthesis. Following ligand binding, some RTKs transduce signals through production of the plasma membrane lipid phosphatidylinositol 3,4,5-triphosphate (PIP3) by phosphoinositide 3-kinase (PI3K). PIP3 acts as an anchor for phosphoinositide-dependent kinase I (PDK1), which will in turn phosphorylate and activate the serine/threonine kinase AKT. Direct targets of AKT are numerous and include transcription factors or repressors such as Foxo3 [95]. An important antagonist of PI3K/AKT signaling is the phosphatase and tensin homolog (PTEN), which functions to dephosphorylate PIP3 to phophatydylinositol 4,5-biphosphate (PIP2), thus abrogating the anchoring of PDK1 to the plasma membrane. FOXO3 is a transcriptional repressor that is now accepted to function as a brake for primordial follicle

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FIG. 3 The balance of activating and maintenance cues in the primordial follicle pool. (A) PI3K/AKT and mTORC pathway activation, which promotes the exit of primordial follicles from quiescence, is restrained by molecules such as Pten and the Tsc1/2 complex. The transcriptional repressor FOXO3 also serves as a molecular brake for primordial follicle recruitment. Growing follicles secrete AMH that acts as a maintenance signal for the primordial follicle pool, most likely acting within the granulosa cells of these primordial follicles. Balanced activation and maintenance signals result in cyclic recruitment of primordial follicles throughout the reproductive lifespan. (B) Genetic models that abrogate the function of maintenance molecules lead to unrestrained activation of the PI3K/AKT/mTORC pathway. This results in activation of the entire primordial follicle pool that leads to exhaustion of the follicular reserve and premature reproductive senescence. (C) Conversely, genetic models that prevent activation of the PI3K/AKT/ mTORC pathway lack follicular development beyond the primordial stage. In these models, a permanent block in primordial follicle activation will eventually lead to atresia and premature reproductive senescence.

activation. Initial studies revealed premature reproductive senescence due to unrestricted, global activation of the primordial follicle pool in a Foxo3-/- mouse line (Fig. 3) [96]. In wild-type ovaries, FOXO3 is localized to both nucleus and cytoplasm of oocytes of primordial and primary follicles [97]. FOXO3 then becomes undetectable in oocytes of secondary follicles and beyond [97]. Conditional Foxo3 ablation from the oocyte [98] leads to global primordial follicle activation identical to that observed in the conventional Foxo3-/- ovary, confirming its oocyte-centric function [99]. Identification of FOXO3 as a likely suppressor of oocyte activation prompted follow-up investigations into the PI3K/AKT pathway as its potential upstream regulator. Conditional deletion of the negative regulator of the PI3K/AKT pathway, Pten, from the oocyte leads to activation of the entire primordial follicle pool, resulting in its premature depletion and ovarian failure by early adulthood [100]. In a similar model of Pten knockout from

oocytes, generated using a distinct recombination strategy, it was shown that the resulting unrestrained activation of the PI3K/AKT pathway leads to hyperphosphorylation, and thus degradation, of FOXO3 [99]. Interestingly, abrogation of AKT activation through conditional deletion of Pdk1 in the oocyte also leads to infertility [101]. In this mouse model, long-term blockade of primordial follicle growth proved to be detrimental, eventually resulting in clearance of these nongrowing follicles and depletion of the primordial follicle pool by early adulthood [101]. These studies show that AKT activation in the oocyte is both necessary and sufficient for activation of the primordial follicle. Somewhat counterintuitively, however, oocyte-specific constitutive activation of PI3K (PI3KCA), which leads to hyperactivation of AKT, does not completely phenocopy the loss of Pten from the oocyte [102]. By PND50, the primordial follicle pool is not completely depleted in oocyte-specific Pi3kca transgenic mice, yet still shows a reduction by about 50%

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FIG. 4 Intraoocyte signaling pathways and regulation of granulosa cell differentiation in primordial follicle activation. Through several genetic models, activation of the PI3K/AKT and mTOR signaling pathways has been shown to promote the exit of the primordial oocyte from quiescence. The initiating extracellular cues for these pathways still require further investigation, with RTKs such as cKIT as potential candidates. Granulosa cells of primordial follicles likely provide such oocyte-activating factor(s). mTOR and the JAK/STAT3 pathways have been shown to promote pregranulosa cell activation, while the transcription factors FOXL2 and GATA4/6 are known regulators of granulosa cell differentiation. Activation of the PI3K/AKT cascade leads to phosphorylation and inactivation of the transcriptional repressor FOXO3. This allows for expression of genes that promote follicle activation and growth. AKT activation also inhibits the Tsc1/2 complex, which allows for activation of mTORC resulting in the promotion of translation of proteins regulating oocyte growth.

compared to wild-type littermates [102]. The slower rate of global primordial follicle activation in oocyte-specific Pi3kca transgenic mice may be attributed to the increased survival rate of follicles at all stages of development [102], suggesting a secondary role of AKT in maintaining oocyte and follicle survival. Findings regarding the involvement of the PI3K/AKT pathway in primordial follicle activation have led to explorations into the potential use of pharmacological modulation of PI3K/AKT in clinical settings. Increasingly improved outcomes of pediatric cancer cases have led to growing interest in the development of fertilitypreserving regiment against gonadotoxic chemotherapeutics. At the forefront is the cryopreservation of cortical ovarian tissues consisting mostly of dormant primordial follicles, followed by in vitro maturation of follicles and eventually the recovery of fertilizable oocytes [103,104]. Efficient means of promoting activation of cryopreserved primordial follicles may greatly improve fertility outcomes for these cancer survivors. Acute exposure of

neonatal mouse ovaries to a PTEN inhibitor, bpV(pic), resulted in activation of the PI3K/AKT pathway and FOXO3 exclusion in oocytes of primordial follicles [105]. Transplantation of treated ovaries into ovariectomized hosts revealed an enhanced rate of follicular growth as compared to controls and in vitro fertilization of the recovered matured oocytes, followed by embryo transfer, is able to generate viable and fertile offspring [105,106]. Similarly, treatment of human ovarian cortical tissues with bpV(pic) followed by xenograft to immunedeficient mice allows for an increased rate of follicular development to the preovulatory stage and recovery of meiotically competent oocytes [105]. Earlier, we discussed the role of KITL and the c-Kit receptor in regulation of germ cell nest breakdown. Early observations in ovaries of a naturally occurring mouse line with a mutation (Steel Panda) that lead to severely reduced expression of KITL (Kitlg) revealed a block in follicle development at the primordial and primary stages [107]. Abrogation of KITL binding to c-KIT using

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antibody ACK2 was shown to block resumption of growth of primordial follicles in both in vivo and ex vivo models across several different species [26,108,109]. In turn, addition of KITL to ex vivo cultured ovaries promotes primordial follicle activation [109]. c-KIT is a RTK that is localized to the oocyte, and c-KIT inhibition appears to phenocopy the block to primordial follicle activation seen in models of PI3K/AKT inhibition [101]. This suggests that c-KIT activation may be at least partially responsible for PI3K/AKT activation within the oocyte during the primordial to primary follicle transition. Indeed, short-term culture of oocytes that are collected from mouse primordial follicles with KITL leads to nuclear export, and as a consequent inactivation, of the transcriptional repressor FOXO3, which is dependent upon activation of the PI3K/AKT-signaling pathway [110]. An oocyte-specific Kit gain-of-function mutation leads to global primordial follicle activation, while oocyte-specific Kit inactivation leads to an extended block to follicular development at the primordial stage that eventually results in global follicular atresia (Fig. 3) [111]. Hyperactivation of AKT and cytoplasmic localization of Foxo3 are observed in oocyte-specific Kit gain-offunction ovaries, while persistent nuclear localization of Foxo3 is observed in oocyte-specific Kit-inactivated ovaries [111]. While these studies provide strong evidence for Kit involvement in primordial follicle activation, the fact remains that only a limited number of primordial follicles are activated at any given time, regardless of c-Kit expression in the oocyte population [112]. Therefore, it is highly likely that additional regulators, perhaps spatially or temporally modulated cofactors to the c-KIT signaling pathway, are involved in the process of primordial follicle activation. Contribution of mTOR Signaling to Primordial Follicle Activation In addition to inactivation of Foxo3, the PI3K/AKT pathway in the oocyte has also been linked to mammalian target of rapamycin (mTOR) signaling in the regulation of primordial follicle activation. The mTOR pathway (Fig. 4) serves as a regulator of cell metabolism, growth, proliferation, and survival by integrating diverse extracellular and intracellular cues. Central to mTOR signaling is the assembly of the large multidomain mTOR complexes (mTORC1 and mTORC2) of serine-threonine kinases. Instead of induction of gene transcription per se, mTOR signaling promotes protein synthesis through activation of components of the translation machinery following phosphorylation by mTORC. Some targets of mTORC include the eukaryotic initiation factor 4E-binding protein 1 (4E-BP1) and the p70 ribosomal s6 kinase 1 (S6K1). The ribosomal protein S6 (rpS6) is a substrate of S6K1 and its phosphorylation; thus, rpS6 phosphorylation is often used as an experimental indicator of mTOR pathway activation. The activated PI3K/AKT

pathway positively regulates mTORC1 activity through inactivation of the mTORC1 inhibitors TSC1 (tuberous sclerosis complex 1) and TSC2. Characterization of one of the Pten oocyte-specific knockout mouse lines revealed increased phosphorylation of rpS6 as compared to wild-type littermates, implicating the involvement of mTOR signaling in the observed global activation of primordial follicles (Fig. 3) [100]. Inactivation of mTORC1 through treatment with rapamycin is able to partially preserve the primordial follicle pool in the oocyte-specific Ptenknockout ovary, suggesting that functions of PTEN in the oocyte are partially mediated through mTORC1 [113]. Additional evidence for the importance of the mTOR pathway in primordial follicle activation comes from conditional deletion of Tsc1 (OoTsc1
/
) or Tsc2 (OoTsc2
/
) in oocytes resulting in global primordial follicle activation (Fig. 3) [114,115]. S6K1 and rpS6 are hyperphosphorylated in OoTsc1
/
or OoTsc2
/
oocytes, indicating increased activation of the mTOR pathway [114,115]. Global primordial follicle activation in OoTsc1
/
ovaries can be attributed entirely to over activation of mTORC1, since long-term treatment with rapamycin completely abrogates the observed depletion of the primordial follicle pool [114]. While mTORC1 in the oocytes appears to be sufficient for activating primordial follicles, it does not seem to be necessary, due to the compensatory potential of the PI3K/AKT pathway. Regulatory-associated protein of mTORC1 (RPTOR) is a part of the mTORC1 complex and is required for mTORC1 assembly. Conditional deletion of Rptor from the oocyte (OoRptor
/
) results in no reproductive phenotypes despite the loss of mTORC1 activity [116]. AKT is hyperphosphorylated in OoRptor
/
oocytes, suggesting that AKT signaling without mTORC1 activity, presumably through inactivation of FOXO3, is able to support a normal rate of primordial follicle activation [116]. Interestingly, mTOR also appears to be important in maintaining primordial follicle quiescence through signaling within the granulosa cells. Granulosa cell-specific disruption of Rptor (PfGC-Rptor
/
) leads to a prevalent block in follicular activation, resulting in very few growing follicles by juvenile age (Fig. 3) [117]. In contrast, granulosa cell knockout of the mTORC1 negative regulator Tsc1 (PfGC-Tsc1
/
) leads to global activation of primordial follicles, a phenotype that can be reversed by a regimen of rapamycin treatment [117]. Granulosa cells of PfGC-Tsc1
/
ovaries express elevated levels of KITL, which leads to hyperactivation of the c-KIT-mediated PI3K/AKT pathway within oocytes [117]. Indeed, pharmacological abrogation of the PI3K/AKT pathway is able to rescue the global primordial follicle activation phenotype of PfGC-Tsc1
/
ovaries [117], consistent with findings that have been discussed earlier in this section. This result suggests that granulosa cell-secreted

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factors, such as KITL, likely act as initiating cues for the oocyte to exit dormancy and for the follicle to enter the growing pool (Fig. 4).

Oocyte-Specific Transcriptional Networks That Promote Primordial Follicle Activation Several oocyte-specific transcription factors are important in the transition from the primordial to the primary follicle stage. The search for master regulators of primordial follicle activation has been performed by a combination of in silico analysis, gene expression studies, and genetic mouse models. Nobox is one of the first oocytespecific homeobox genes identified through in silico analysis of NCBI UniGene database of a neonatal mouse ovary cDNA library [118]. Detected in oocytes beginning at E15.5, high expression of Nobox remains throughout folliculogenesis. While the males are completely fertile, Nobox
/
females are infertile due to an almost complete loss of oocytes by 6 weeks of age [119]. Although primordial follicles are initially observed, no growing follicles are ever detected in Nobox
/
ovaries, suggesting a failure in primordial follicle activation. By PND14, Nobox
/
ovaries contain mostly degenerating follicles which lack oocytes. At the molecular level, Nobox binds to the promoters of the oocyte specific genes Oct4 and Gdf9, and expression of these genes is lost in Nobox
/
ovaries [119,120]. This finding is consistent with the clinical observation of high prevalence of NOBOX mutations, leading to compromised ability to bind the Gdf9 promoter, in patients with primary ovarian insufficiency [121]. The lim-homeobox protein, LHX8 is localized to oocytes throughout follicle development [122]. While conventional knockout of Lhx8 results in very similar phenotypes to those observed in Nobox
/
mice, characterized by failure of primordial follicles to transition into the growing pool and eventual global atresia by PND7, conditional oocyte deletion of Lhx8 reveals a more specific function [123], with global primordial follicle activation due to hyperactivation of the PI3K/AKT pathway [124]. LIN28A, a known positive regulator of the AKT/ mTOR pathway, is elevated in the oocyte-specific Lhx8 knockout, and indeed, Lhx8 was found to bind the Lin28a promoter and suppress its expression [124]. Sohlh1 and Sohlh2 encode basic helix-loop-helix transcription factors that are expressed exclusively in female germ cells beginning at E15.5 and E12.5, respectively [122,125,126]. While expression of Sohlh1 and Sohlh2 is robust in oocytes within germ cell nests and primordial follicles, their expression diminishes in primary follicles and beyond, suggesting that their functions are restricted to early follicular development. Sohlh1 [122] or Sohlh2 [127] knockout females are infertile due to lack of follicular growth. While the size of the primordial follicle pool in

33

Sohlh1
/
and Sohlh2
/
ovaries is comparable to wildtype littermates, follicles with cuboidal granulosa cells are never observed, indicating a failure of primordial follicles to transition into the primary stage [122,127]. SOHLH1 and SOHLH2 homodimerize, as well as heterodimerize, and promote expression of oocyte specific genes such as Nobox and Lhx8 [126,128]. Collectively, these studies begin to clarify a transcriptional network that regulates the process whereby primordial follicles remain dormant, or are activated to form primary follicles.

Granulosa Cell Differentiation During the Primordial to Primary Follicle Transition As discussed in the last section, granulosa cells of primordial follicle likely provide ligands or extracellular cues that initiate the intra-oocyte cascades that lead to follicle activation. Recently, activation of the JAK/STAT3 pathway in granulosa cells of primordial follicles was shown to enhance primordial follicle recruitment [129]. Further analysis on the downstream consequences, and particularly of the changes in secreted products following JAK/STAT3 pathway activation, may aid in identifying additional candidates for primordial oocyte-activating factors. Failure of granulosa cells to undergo proper differentiation often leads to unsynchronized development between the oocyte and granulosa cells, which is detrimental to the survival of the follicle. Previously, we discussed the failure of pregranulosa cells to interact with germ cell nests and properly form follicles in Foxl2
/
mice [130]. A distinct transgenic line of Foxl2 disruption (Foxl2lacZ), while appearing to undergo primordial follicle formation, is infertile due to a block in follicular development during the primordial to primary stage transition [23]. Foxl2lacZ squamous granulosa cells never become cuboidal, and thus remain nonproliferative [23,85]. Interestingly, in 2-week-old Foxl2lacZ ovaries, Kitl expression is significantly upregulated in the granulosa cells, resulting in stimulation of Gdf9 expression in almost all oocytes, a hallmark of a growing oocyte population, and thus global follicular activation [131]. Despite the failure of the granulosa cells to differentiate from the dormant squamous morphology, oocytes do undergo activation in Foxl2lacZ ovaries, suggesting a failure of the germ cells and somatic cells to coordinate their developmental transitions [23]. The importance of proper differentiation of granulosa cells to follicle survival and primordial to primary stage transition is further illustrated by a transgenic mouse targeting the gonadal specific transcription factors GATA4 and GATA6. GATA proteins are zinc finger transcriptions factors, and GATA4 and GATA6 are predominantly expressed in granulosa cells beginning at the embryonic age in the mammalian ovary and function to regulate

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expression of numerous genes that are important throughout follicular development, including those involved in follicular growth and steroid biosynthesis [132,133]. Deletion of both Gata4 and Gata6 in granulosa cells leads to a loss of expression of granulosa cell markers, including Foxl2, indicating compromised granulosa cell identity [134]. Consistent with the most prominent phenotype of Foxl2lacZ ovaries, granulosa cells do not transition from a squamous to cuboidal morphology following Gata4 and Gata6 deletion, leading to a lack of growing follicles and eventually to clearance of the permanently dormant primordial follicle pool through wide-spread atresia [134]. While currently less well understood than the expansive oocyte-specific transcriptional network that is implicated in primordial follicle activation, future investigations into novel FOXL2 and GATA4/6 targets should provide further important insights into the concurrent differentiation of follicular granulosa cells.

CELLULAR SIGNALING DURING PREANTRAL FOLLICULAR DEVELOPMENT The growth of follicles between the primary and antral stages is accepted to be independent from the pituitary gonadotropin, FSH. This paradigm is derived from the unaltered preantral follicular development observed in

various models of FSH signaling disruption [135–137]. While gonadotropin-responsive antral follicles are never observed in genetic models lacking FSH (through abrogation of the β-subunit of FSH) [135] or lacking the FSH receptor (FSHR) [136,137], the numbers of primordial, primary, and multilayered preantral follicles are unaffected. This suggests that local signals are predominant in regulating the development of follicles through the preantral stages. The acquisition of additional layers of granulosa cells during the transition between the primary and the secondary follicular stages and beyond involves many diverse, and often highly interacting signaling pathways. Moreover, during this period, the oocytes undergo size expansion that needs to be tightly coordinated with the growth of the granulosa cells. Finally, as follicles reach the secondary stage, local signals are required to initiate the recruitment and organization of the thecal cell layer, a second type of somatic cell that is indispensable for steroidogenesis. Fig. 5 illustrates the diversity of signaling pathways that are involved in gonadotropinindependent granulosa cell growth, as well as the multidirectional communication between the oocyte and somatic cells during this preantral period. While we recognize the complexity of many pathways involved in these preantral developmental events, we first focus discussion on paracrine signaling through the TGF-β superfamily of peptides, whose members are highly represented among both oocyte-secreted and somatic FIG. 5 Regulation of gonadotropinindependent follicular growth. The growth of ovarian follicles prior to antrum formation is independent of FSH and is governed by various modes of local signaling. Following activation, oocytes begin secreting GDF9 and BMP15, which act on granulosa cells to promote growth and gene expression. Under the influence of GDF9, granulosa cells express the Hh ligand that functions to specify the final somatic cell layer, the theca cells. Activin is a mitogen that is secreted and signals within the granulosa cell layer. Activin promotes expression of Esr2 and suppresses expression of Cyp26b1. This results in increased levels of ERβ signaling and retinoic acid receptor signaling, both of which are growth promoting within granulosa cells. Promotion of granulosa cell proliferation is also achieved through cross-regulation between the activin and Notch signaling pathways. Notch signaling occurs between the oocyte and granulosa cells, as well as between adjacent granulosa cells.

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cell-secreted factors in the ovary (reviewed in Ref. [71]). In addition, we discuss the role of the juxtacrine Notch signaling pathway that is important for preantral follicular growth, in addition to its roles during germ cell nest breakdown. Finally, we summarize recent findings on the functions of the highly conserved Hedgehog signaling pathway in regulating the formation of the thecal cells.

TGF-β Family Signal Transduction in the Ovary Based on ligand sequence similarity and modes of signal transduction, the TGF-β superfamily is divided into two subgroups, the growth differentiation factors/bone morphogenetic proteins (GDFs/BMPs) and the activins/TGF-βs. TGF-β superfamily peptides form either homo- or hetero-dimeric ligands. Activins and TGF-βs initiate signaling by binding to a serine/threonine kinase cell surface receptor, known as the type II receptor [138]. Five type-II receptors have been identified in mammals [139]. The type-II receptor recruits a type-I receptor, also known as an activin-like kinase (ALK), and activates the type-1 receptor through trans-phosphorylation [138]. Seven type-1 receptors have been characterized in mammals [139]. In contrast, BMPs bind first to a type-I receptor, followed by recruitment of a type-II receptor, or to a preformed type-1/type-2 receptor complex on the cell surface [140]. The signal is transduced intracellularly by type-1 receptor-mediated phosphorylation of a SMAD protein (receptor-regulated SMAD -rSMAD), which will then recruit an obligatory common-mediator SMAD (coSMAD) [141]. Receptor activation by GDFs/BMPs activates SMAD1, SMAD5, or SMAD8, while receptor activation by activins or TGF-β activates SMAD2 or SMAD3 [142]. One exception to this model is the ligand GDF9, which utilizes SMAD2/3 in its signal transduction [143,144] SMAD4 acts as a coSMAD for receptor activation by both subtypes of ligands [142]. Finally, an rSMAD-coSMAD complex translocates to the nucleus and acts as a transcriptional activator of target genes [141]. The SMAD pathway is negatively regulated by inhibitory SMADs (SMAD6, SMAD7) through prevention of rSMAD activation at the receptor level or through competitive binding of activated rSMAD to coSMAD [141]. In granulosa cells, there appears to be significant functional redundancies between SMAD1, 5, and 8, and between SMAD2 and 3. This is demonstrated by the absence of fertility defects following disruption of any of these individual rSmad alleles in female mice [145,146]. Since conventional knockouts of all but Smad8 result in embryonic lethality, a conditional knockout approach has been taken to study these pathways in the ovary. Double conditional knockout of Smad1/5 or triple conditional knockout of Smad1/5/8 in granulosa cells

35

results in infertility due to development of metastatic granulosa cell tumors by 3 months of age [145]. In contrast, the double conditional knockout of Smad2/3 is subfertile, showing decreased numbers of antral follicles and increased follicular atresia as indicated by a high incidence of ZP remnants in 3-month-old ovaries [146]. In addition to follicular developmental aberrations, a high incidence of trapped oocytes in luteinizing follicles and a reduced expression of cumulus expansion markers strongly suggest ovulatory defects in Smad2/3 conditional knockout mice [146]. Granulosa cell conditional knockout of the coSMAD Smad4 results in similar phenotypes to the Smad2/3 conditional knockout, presenting with increased numbers of atretic preantral follicles and ovulatory defects [147]. Control of Follicular Growth by Oocyte-Secreted GDF9 and BMP15 Immediately following activation, oocytes of primary follicles begin to produce GDF9, a secreted TGF-β family ligand [131,148,149]. Gdf9 null female mice are infertile due to a block in follicular development at the primary follicle stage, which appears to result from a failure of granulosa cell proliferation [131,150,151]. Demise of the follicles is initiated by degeneration of the oocytes in Gdf9
/
ovaries, which results in surviving follicular remnants that acquire a luteinized morphology [131]. The actions of GDF9 appear to be specifically directed toward somatic cells, since Gdf9
/
oocytes can be successfully matured in vitro (with addition of the necessary factors) to resume meiosis [150]. In vivo treatment of PND5 rats with GDF9 leads to increased activation of primordial follicles and subsequent enhancement of growth of primary follicles into multilayered preantral follicles [152]. Gdf9 stimulates proliferation in cultured granulosa cells, as well as growth of cultured primary and secondary follicles [153,154]. The actions of GDF9 on granulosa cells are most likely achieved through binding with a complex of the type-I receptor ALK5 [143,155] and the bone morphogenetic protein receptor type II (BMPRII) [156] as disruption of either leads to abrogation of the proliferative effects of GDF9 on granulosa cells [143, 155, 156]. BMP15, a TGF-β family ligand highly related to GDF9, was identified through a homology-based cloning strategy [157]. Like Gdf9, Bmp15 is expressed specifically in oocytes beginning at the primary follicle stage [157] and exerts proliferative effects on cultured rat granulosa cells [158]. Several naturally occurring mutations affecting BMP15 signaling, and as a consequent ovulation rate, have been characterized in sheep. The Inverdale (FecXI) mutation is a point mutation that leads to a single amino acid replacement in the mature Bmp15 protein, while a distinct point mutation known as the Hanna (FecXH) mutation leads to the creation of premature stop codon and thus a truncated Bmp15 protein [159]. Both the FecXI and FecXH alleles lead to nonfunctional Bmp15, and

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homozygous females are infertile due to a block in follicle development at the primary stage [159]. Counterintuitively, ewes heterozygous for either mutation have an increased ovulation rate, which leads to a higher incidence of multiple births compared to wild-type ewes [159]. Consistent with the mitogenic role of Bmp15 in granulosa cell growth, antral follicles in heterozygous FecXI and FecXH ewes are undersized; yet, the granulosa cells of these antral follicles have increased expression of Lhcgr, rendering them more responsive to the LH-ovulatory signal [160]. Similarly, Booroola (FecB) ewes with a mutation that leads to reduced intracellular kinase activity of the type I BMPRIB (ALK6) receptor are hyperprolific, with high rates of multiple births [161]. While the precise molecular basis of the dependency of ovulation rate on the level of BMP15 signaling remains unclear, it has been postulated that the determining factor may be the ratio of GDF9 to BMP15 in the ovary [162]. Given the similarity in the structures of BMP15 and GDF9, as well as their expression by the oocyte in a temporally overlapping manner, it is of interest to understand whether they assert their effects on granulosa cells in any cooperative or redundant manner. When coexpressed in human embryonic kidney 293T (293T) cells, recombinant GDF9 and BMP15 are able to form heterodimers, suggesting their potential interaction in vivo [163,164]. In the mouse model, Bmp15
/
females are subfertile, in contrast to the infertile Gdf9
/
females, presenting with only minimal morphological defects in follicular development, albeit having decreased ovulation and fertilization rates [165]. A more severe fertility defect was observed in Bmp15
/
Gdf9+/
females compared to Bmp15
/
females [165]. However, Bmp15
/
Gdf9
/
double-knockout females have a fertility defect that is no worse than that of the Gdf9
/
single mutant [165]. In cultured mouse granulosa cells, GDF9 and BMP15 were shown to act synergistically to induce proliferation and SMAD3-dependent transcription [166]. The importance of GDF9:BMP15 heterodimers in regulating granulosa cell function was further demonstrated in a mouse cumulus cell expansion assay, where they were shown to be 10- to 30-fold more potent than the GDF9 homodimer in promoting cumulus expansion through activation of the SMAD2/3 pathway [167]. Several mutations at the BMP15 locus have been associated with cases of primary ovarian insufficiency (POI) in humans [168]. Biochemical investigations into ten of these POI-associated BMP15 mutations showed that they resulted in reduced levels of mature BMP15, a decreased ability of BMP15 to induce SMAD1/5/8-dependent transcription, or a reduced capacity for BMP15 to synergistically induce proliferation and SMAD3-dependent transcription in the presence of GDF9 [168]. Similarly, mutations in the GDF9 locus have also been identified in cases of POI, including those that lead to aberrations

in the GDF9 transcriptional regulatory region [169] and point mutations that lead to reduced levels of mature GDF9 or to aberrant mature GDF9 that is less potent in activating the SMAD2 pathway [170]. These findings suggest that the cumulative effects of BMP15 and GDF9 on granulosa cell proliferation and gene expression are a complex function of the two distinct rSmad pathways that they activate. Regulation of Granulosa Cell Proliferation by Granulosa Cell-Derived TGF-β Family Ligands In addition to GDF9 and BMP15 originating from the oocyte, granulosa cells also produce multiple TGF-β family ligands that act in a paracrine manner to regulate follicular development. There are three mammalian TGF-β isoforms (TGF-β1–3), and localization studies have identified their expression in all three follicular cell types beginning at the preantral or early-antral stages in human and rodent ovaries [171,172]. Depending on the species, TGF-β1 can assert stimulatory or inhibitory effects on granulosa cell proliferation. While in vitro, TGF-β1 promotes rat granulosa cell proliferation, it exerts inhibitory effect on the proliferation of granulosa cells from cattle, sheep, and pig ovaries (reviewed in Ref. [171]). In contrast, negative regulation of thecal cell steroidogenesis by TGF-β1 appears to be conserved across these species [171]. Activins and inhibins are granulosa cell-secreted peptide hormones that were originally identified as a part of the ovarian feedback loop regulating the release of FSH by the pituitary gland [71]. Mature activin and inhibin are composed of dimers of structurally related peptides connected by a disulfide bond [173]. Dimers of the two β subunits form mature activins (Activin A—βAβA, Activin B—βBβB, or Activin AB—βAβB), while heterodimers of the α and β subunits result in mature inhibins (Inhibin A—αβA or Inhibin B—αβB). Expression of all three subunits can be detected in granulosa cells and is dynamically regulated throughout the rat estrous cycle [48]. Activin exerts its actions through binding to its type-II receptor (ACTRIIA or ACTRIIB) and transactivation of ALK2, 4, or 7, followed by activation of SMAD2/3 and the co-SMAD SMAD4 (reviewed in Ref. [174]). Inhibin suppresses the actions of activin through competitive binding with the type-II receptor. Finally, as previously discussed, the ovary also produces FST, a molecule that binds to and neutralizes the biological activity of activin [175]. In vitro, activin exerts proliferative effects on granulosa cells of preantral follicles isolated from prepubertal mice, and FST treatment blocks this effect [176]. The effects of activin on granulosa cell proliferation appear to be age dependent, possibly due to additional growth signals by FSH following the onset of puberty. For example, activin treatment of neonatal mice leads to increased

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CELLULAR SIGNALING DURING PREANTRAL FOLLICULAR DEVELOPMENT

proliferation of granulosa cells and to a limited degree the germ cells [21]. However, while activin stimulates growth of preantral follicles isolated from juvenile mice, it promotes atresia and abrogates FSH-induced growth in preantral follicles isolated from adult animals [177,178]. Regardless of this age-specific effect, activin appears to be necessary for female fertility, as animals lacking any ovarian expression of either of the β subunits are infertile, showing decreased expression of factors that are associated with granulosa cell survival and proliferation such as Kitl, Taf4b, and c-Myc [179]. Consistent with a deficiency in activin signaling, juvenile Smad3
/
ovaries have decreased numbers of preantral and early antral follicles, indicating disrupted follicular growth from the secondary stage and beyond [180]. Overexpression of the inhibin α subunit in transgenic mice (MT-alpha) results in increased levels of circulating mature inhibins and decreased levels of mature activins [181]. Expression of both β subunits is downregulated in the MT-alpha ovaries, suggesting a certain degree of autoregulation of gene expression by activin signaling [181]. Consistent with pharmacological and genetic models of suppressed activin action discussed above, MT-alpha female mice are subfertile with decreased numbers of mature antral follicles in the ovary [181,182]. While inhibin is not currently known to activate any specific receptor, the genetic model of inhibin deficiency Inha
/
has demonstrated the necessity for its presence in order to restrain and balance the mitogenic effects of activin on granulosa cells [183]. Inha
/
ovaries are abnormally large compared to the wild-type littermates due to granulosa cell hyperproliferation, which eventually leads to gonadal tumors and infertility in adulthood [183]. Studies in cultured granulosa cells from preantral follicles have revealed the potential for cross-talk between activin and at least two different types of nuclear receptor signaling pathways in regulating granulosa cell proliferation (Fig. 5). Activin treatment is found to induce expression of Esr1 and Esr2 in a SMAD2/3-dependent manner in proliferating granulosa cells [184]. Since estrogen is mitogenic, as well as pro-survival, in preantral granulosa cells [185–187], it is likely that activin promotes proliferation at least in part through enhancement of estrogen receptor signaling. Transcriptomic analysis of activin-treated granulosa cells identified Cyp26b1 as downregulated by activin [188]. A member of the cytochrome P450 family, the gene product of Cyp26b1 functions to degrade RA and is expressed in granulosa cells of growing follicles [188]. RA or Cyp26b1 inhibitor treatments of cultured granulosa cells leads to an increase in granulosa cell proliferation in a dose-dependent manner, while inhibition of retinoic acid receptors abrogates this effect [188]. Besides identifying a novel mitogenic factor in

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granulosa cells, this study further illustrates the extensive network of signals with which activin interacts during gonadotropin-independent follicle growth.

The Role of Notch Signaling in Preantral Follicle Development Earlier, we reviewed evidence for a role of Notch signaling in promoting germ cell nest breakdown. It is now appreciated that Notch signaling continues to be important during follicle growth and development, functioning to promote granulosa cell proliferation and to coordinate the development of the oocyte and granulosa cells (Fig. 5). Inhibition of Notch signaling using the γ-secretase inhibitors, DAPT or L658,458, leads to decreased proliferation of cultured granulosa cells or secondary follicles [189]. Expression of c-Myc, a proliferation-associated transcription factor, in granulosa cells is suppressed following Notch inhibition, and restoration of Notch signaling through expression of constitutively active N2ICD leads to recovery of both c-Myc expression and granulosa cell proliferation [189]. Additionally, Notch may also regulate granulosa cell proliferation through interactions with activin signaling. There is evidence that the Notch target gene Hey2 is coregulated by activin in a SMAD3-dependent manner in preantral granulosa cells [190]. Indeed, activin treatment is able to overcome the effect of DAPT inhibition and restore proliferation in cultured granulosa cells [190]. In conditional-knockout models of the ligand JAG1 from the oocytes (J1KO) or the receptor NOTCH2 from granulosa cells (N2KO), defects in proliferation of granulosa cells are first observed by PND19, and are associated with enhanced cell death [74]. J1KO and N2KO ovaries also present with a phenotype that indicates disruption of developmental coordination between oocytes and granulosa cells, characterized by enlarged oocytes that are contained in very early stage growing follicles (i.e., follicles with a single layer of granulosa cells) [74]. Molecularly, this is supported by elevated expression of the oocyte-specific factors Gdf9, Figla, and Zp3 as compared to wild-type littermates [74]. The Notch target gene Hes1 is expressed in somatic cells embryonically (by E15.5) and its expression decreases following birth [71,191]. Conventional, or granulosa cell conditional, knockouts of Hes1 lead not only to increased oocyte death but also increased proliferation of the pregranulosa cells [191]. Both mouse lines are subfertile, indicating that disruption of early follicular development results in detrimental effects on fecundity. Increased expression of the βB subunit (Inhbb) of activin is observed in both models of Hes1 deficiency [191], which makes the enhanced pregranulosa cell proliferation consistent with effects observed in neonatal mice

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treated with exogenous activin [21]. An additional common phenotype between the two models of Hes1 deficiency is the suppression of c-Kit expression in oocytes [191]. As covered in earlier sections, KITL/c-KIT signaling is particularly important for direct communication between granulosa cells and oocytes, and it regulates nest breakdown and primordial follicle activation in the neonatal ovary. Thus, while Notch signaling is mitogenic in preantral granulosa cells, an additional function is perhaps to ensure coordinated development between oocytes and granulosa cells prior to exposure to gonadotropins, which will act as the master regulators for follicular development beyond the preantral stage.

Molecular Mechanisms of Theca Cell Recruitment and Specification The thecal cell layer forms just outside the basement membrane surrounding the outermost granulosa cells when the follicle reaches the secondary stage [192]. Through the use of transgenic reporter mouse lines and lineage tracing, it was shown that theca cells originate from one of two sources, the fibroblast-like precursor cells indigenous to the ovary (Wt1 expressing cells) or mesenchymal cells that migrate into the ovary from the mesonephros [193,194]. Theca cells function in ovarian steroidogenesis by providing aromatizable androstenedione to the granulosa cells, where aromatase will convert the androgen into 17β-estradiol [192]. It was long suspected that preantral follicles secrete certain thecal cell differentiating factors, as conditioned medium collected from theca-less preantral follicles stimulated androstenedione production in cultured thecal-interstitial cells [195]. This putative theca differentiating factor was thought to be a small molecule on the order of 20–25 kDa in size [196]. Recent studies indicate that the developmentally conserved Hedgehog (Hh) signaling pathway is the chief regulator of thecal cell specification in the ovary. Hedgehog signaling is initiated by binding of one of three HH ligands (Sonic hedgehog—SHH, Indian hedgehog— IHH, Desert hedgehog—DHH) to the canonical receptor Patched (PTCH1, 2). Hh binding to PTCH results in derepression of Smoothened (SMO2), an obligate signal transducer in Hh signaling, and finally the translocation of the Hh transcriptional effector GLI (GLI1 in thecal cell precursors) into the nucleus. Ihh and Dhh are expressed by granulosa cells of growing follicles, while the receptors Ptch1, Ptch2, and the transcriptional effector Gli1 are only detected in the surrounding pretheca cells, suggesting that theca cells are the exclusive target of Hh ligands in the ovary (Fig. 5) [197]. The thecal layer never forms in ovaries with conditional deletion of both Dhh and Ihh from the granulosa cells [194]. These animals are infertile, with blocked

follicular development at the preantral stage and blunted steroid production [194]. It has been shown that GDF9 from the oocyte stimulates the expression of Dhh and Ihh in granulosa cells, and indeed, expression of both Hh ligands is severely downregulated in Gdf9
/
ovaries [194]. Theca layer recruitment and differentiation is an elegant example of how three distinct cell types within the follicle communicate and regulate each other’s functions through diverse signaling pathways.

REGULATION OF GRANULOSA CELL PROLIFERATION AND DIFFERENTIATION IN ANTRAL AND PREOVULATORY FOLLICLES In larger multilayer follicles, small fluid filled spaces are initiated that will eventually converge to form the antral cavity [198]. A hallmark of antral follicle development is the increased abundance of transcripts for the FSH receptor (Fshr) within the granulosa cells, which upon translation into FSHR renders the follicles responsive to circulating FSH [199]. Indeed, FSHR signaling is required for continued development beyond the multilayered secondary follicle stage, as antral follicles are never detected in mice lacking either mature FSH (Fshb
/
line) [135] or FSHR [136,137]. Following puberty, FSH functions to prevent early antral follicles from undergoing atresia [200] by regulating both granulosa cell proliferation and differentiation [201,202]. Under the influence of FSH, the cell cycle of granulosa cells is shortened, requiring a doubling period of only about 24 h as compared to requiring greater than 7 d during the FSHindependent period [203]. FSH also functions to induce gene expression changes in granulosa cells, which differentiates them toward the LH responsive, preovulatory stage [204,205]. Characteristics of granulosa cell maturation in response to FSH signaling include the expression of genes involved in steroid biosynthesis, such as Cyp19a1, Cyp11a1, and Hsd3b1, which leads to increased production of estradiol and progesterone, and the expression of the membrane receptor for LH (Lhcgr) [204,206]. The FSHR is a G-protein-coupled receptor (GPCR) that stimulates synthesis of the second messenger cyclic AMP (cAMP) when activated [207,208]. The intracellular rise in cAMP leads to activation of the cAMP-dependent protein kinase A (PKA) [209], which in turn promotes gene transcription through phosphorylation and activation of several transcription factors, including but not limited to, the cAMP response element-binding protein (CREB) [210,211] and GATA4 [212,213]. Additionally, PKA downstream of FSHR activation also modifies chromatin structure through phosphorylation of histone H3, which leads to increased nucleosome accessibility [214,215]. It is now appreciated that to exert maximum mitogenic and differentiation effects, FSH/FSHR signaling interacts

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and cooperates with diverse local ovarian signals. In this section, we discuss intraovarian signals that regulate proliferation and differentiation in antral to preovulatory granulosa cells and their cross talk with FSHR signaling. For the purposes of clarity and simplicity, we will refer to granulosa cells as undifferentiated prior exposure to FSH (including those collected from estrogen stimulated animals and used in vitro studies) and as differentiating following activation of FSHR, while appreciating that these are points along a continuum [216].

Crosstalk Between FSHR Signaling and Kinase Cascades in Granulosa Cell Proliferation and Differentiation Control of Granulosa Cell Proliferation and Gene Expression by MAPK/ERK Activation Downstream of FSHR Signaling The mitogen-activated protein kinases (MAPKs) are a highly conserved family of serine/threonine protein kinases that function to relay extracellular cues by way of kinase cascade activation and influence diverse cellular

39

processes, including proliferation and gene transcription (Fig. 6) [217]. While originally described as an intracellular signaling pathway that is downstream of RTKs, the MAPK pathway is now also known to function downstream of GPCR activation [218]. The MAPK/ERK pathway is active in proliferating cells, including granulosa cells. ERK1/2 activation is observed in undifferentiated proliferating granulosa cells in culture [219]. An increase in intracellular cAMP levels was found to prevent activation of ERK2 in response to growth factors in human arterial smooth muscle cells, adipocytes, and mouse fibroblast cell lines [220]. Based on these findings, FSHR activation and its associated rise in cAMP levels were initially hypothesized to downregulate ERK1/2 activation in differentiating granulosa cells. On the contrary, FSH stimulation of rat granulosa cells in culture leads to an acute activation of ERK1/2, which occurs in a cAMPdependent manner [219]. In addition to proliferation, activation of the MAPK/ERK pathway following FSH stimulation is also important for transcription of genes associated with the preovulatory phenotype, including positive regulation of the Cyp19a1, Cyp11a1, Inha, Lhcgr, and Egfr genes [221].

FIG. 6 Integration of FSH and paracrine signals in promoting granulosa cell proliferation and differentiation. Following antrum formation, granulosa cells proliferate and differentiate into the preovulatory stage through upregulation of Lhcgr and genes important for steroidogenesis in response to FSH. To achieve the maximal proliferative and differentiation effects of FSH, FSHR signaling interacts with kinase cascades downstream of RTKs such as EGFR and IGFR and cooperates with Smad-dependent signaling pathways downstream of TGFβ and activin family receptors. Integration of PKA and PI3K/AKT signaling leads to inactivation of the transcriptional repressor FOXO1 through phosphorylation, which targets it for degradation. In turn, MAPK/ERK and SMAD2/3 activation provide activating signals for the transcription of granulosa cell differentiationassociated genes. Notch signaling promotes granulosa cell differentiation through positive regulation of steroid biosynthetic enzyme gene expression and through suppression of proliferative kinase cascades. Additionally, the Notch NICD is known to coregulate expression of common effectors by interacting with the SMAD complex. Note that distinct subsets of transcriptional regulatory factors will interact at the promoters of different target genes, and the schematic illustrates generic and summed examples of such regulatory pathway interactions. I. THE OVARIAN FOLLICULAR APPARATUS: OPERATIONAL CHARACTERISTICS

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It is still debated whether any specific RTK is involved in MAPK/ERK pathway activation in FSH-stimulated granulosa cells. In undifferentiated, proliferating granulosa cells from estrogen-treated mice, the MAPK pathway upstream of ERK1/2 is observed to be constitutively active [222]. In these granulosa cells, ERK1/2 is continuously dephosphorylated by the MAP kinase phosphatase 3 (MKP3), thus restraining the full activation of the MAPK/ERK pathway [221]. FSH stimulation and PKA activation were in turn shown to inhibit MKP3 activity, and thus function to relieve the inhibition on ERK1/2 [221]. Gene expression changes are in turn dependent upon phosphorylation of the RNA and DNA binding protein YB-1 by the activated ERK1/2 [220]. While epidermal growth factor receptor (EGFR) functions prominently in propagating the LH signal during ovulation, there is also evidence for its modulation downstream of FSHR signaling. Radioactively labeled epidermal growth factor (EGF) is observed to bind to immature rat follicles, and FSH treatment enhances the number of EGF molecules bound, suggesting modulation of Egfr expression by FSH in differentiating granulosa cells [223]. More recently, Egfr expression was shown to be strongly downregulated in Fshb
/
prepubertal ovaries [224]. Treatment with FSH was shown to restore Egfr expression (and eventually the ability to ovulate), confirming modulation of Egfr by FSHR signaling. In addition to MAPK/ERK activation downstream of FSHR, a known, but minor, product of Fshr alternative splicing (growth factor type1 receptor or oFSH-R3) whose activation rapidly leads to phosphorylation of ERK1/2 instead of the rise in cAMP level can also be detected in the ovary [225]. Through the use of the TNR mouse line, Notch signaling is observed to remain active in granulosa cells of antral and preovulatory follicles [226]. Notch signaling molecules are expressed in these later stage follicles and there is evidence that their expression is dynamically regulated by gonadotropins [72]. Of note, is the ligand Jag1 whose localization shifts from being oocyte-specific to being present in ovarian somatic cells following exposure to exogenous gonadotropins [226,227]. Disruption of the Notch ligand Jag1 in cultured granulosa cells collected from PMSG-primed immature mice leads to a compromised differentiation response, as measured by suppressed steroidogenesis [226]. At the expense of suppressed differentiation, enhanced proliferation, resembling that of less mature granulosa cells, is maintained in Jag1-deficient cells and this is associated with to increased activation of the MAPK/ERK pathway. Together, these studies demonstrate that in differentiating granulosa cells, FSH functions to promote proliferation and differentiation in conjunction with underlying signals that are initiated by local factors. Regulation of Granulosa Cell Differentiation and Proliferation by the PI3K/AKT Pathway Interest in insulin growth factor receptor (IGFR), a RTK, signaling in follicle maturation began with the

observation that the ligand insulin-like growth factor-1 (Igf1) and Fshr are coexpressed in healthy, steroidogenically active large antral follicles from mouse [228]. Igf1 null females are infertile, with ovaries lacking follicles beyond the early antral stage, resembling Fshb
/
animals [228]. In granulosa cells, activation of the IGF1-R or the FSHR independently stimulates the PI3K/AKT pathway (Fig. 6), and coactivation of both receptors leads to synergistic effects on AKT activation [229]. Indeed, inhibition of the IGF1-R prevents the induction of granulosa cell differentiation by FSH, as measured by the lack of an increase in Cyp19a1 expression and estradiol production [229]. Additional support for the importance of IGF1-R signaling in facilitating follicular maturation by FSH comes from the recently reported Igf1r granulosa cell conditional knockout (IGF1Rgcko) mouse line [230]. IGF1Rgcko females are infertile with ovaries lacking antral follicles and granulosa cells that are unable to differentiate in response to pregnant mare serum gonadotropin (PMSG) stimulation, which activates the FSHR, despite normal levels of Fshr expression compared to wild-type littermates [230]. Another important interaction between FSHR signaling and a paracrine factor occurs in the regulation of granulosa cell proliferation by activin. In cultured rat granulosa cells, the full mitogenic effect of FSH requires the presence of activin [231]. The apparent explanation is that activation of both AKT and SMAD2/3 is required for expression of the cell cycle regulator Cyclin D2 (Ccnd2) following FSH and activin stimulation (Fig. 6) [231]. It is now appreciated that FSH activation of the PI3K/AKT pathway occurs through a PKA-dependent cascade [232] and targets the transcriptional suppressor Foxo1 [233]. Phosphorylation of Foxo1 leads to its translocation from the nucleus and directs it for degradation. More than 60% of FSH-regulated genes have now been identified to be targets for Foxo1, including those involved in proliferation, such as Ccnd2, and those involved in steroid biosynthesis, such as Star, Cyp11a1, and Cyp19a1 [233,234]. Fig. 6 summarizes the integration of FSHR, RTKs, Notch, and activin signaling pathways in regulating granulosa cell proliferation and differentiation. These data demonstrate the importance of, what are often accepted to be ubiquitous, kinase cascades in integrating endocrine and paracrine cues to promote the differentiated preovulatory phenotype in granulosa cells.

The Roles of Steroid Hormones in Follicular Maturation The ovarian follicle is the main source of sex steroid hormones (estrogens, progestins, and androgens) that are indispensable for overall female reproductive physiology. Sex steroids act on many organ systems, including but not limited to the nervous system, the skeletal system, the cardiovascular system, metabolism, and last but not

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least the reproductive system. Comprehensive discussions of the molecular mechanism of ovarian steroidogenesis are found elsewhere in this text and reviewed in Ref. [235]. The two-cell, two-gonadotropin model of follicular steroidogenesis [236] is based on localization of the enzymes required for distinct steps in the steroid biosynthetic cascade. Theca cells convert cholesterol into androstenedione under the influence of Lhcgr signaling. In turn, the hydrolyzable androstenedione is utilized by granulosa cells as a substrate for estradiol under the influence of FSHR signaling. Following ovulation and formation of the corpus luteum, progesterone will become the main sex steroid produced by the ovary. In this section, our focus will be on the role of estrogens and androgens as paracrine factors regulating follicular events throughout the FSH-dependent and periovulatory-developmental stages. While we recognize the equally important role of progesterone, we refer readers to Ref. [237] for a discussion on the importance of progesterone receptor signaling during the ovulatory cascade and pregnancy. Estrogen and Estrogen Receptor Signaling in Antral and Preovulatory Follicles Estrogen increases granulosa cell proliferation in preantral follicles [185–187]. Estrogen is also found to increase the presence of FSHR and LHCGR in granulosa cells and enhances the rise in cAMP following cholera toxin stimulation (due to blocked hydrolysis of G-protein-associated GTP, and thus constitutively activated adenylate cyclase), suggesting its ability to augment GPCR/cAMP signaling [238]. Ovaries of mice lacking aromatase (Cyp19a1; ArKO), an enzyme that is required for the final conversion of androgens into estradiol, lack follicles beyond the early antral stages [187]. The phenotypes of the ArKO mouse, however, appear to be caused by chronic elevated serum gonadotropin levels due to the lack of estrogen feedback to the pituitary. Upon exogenous estrogen treatment, follicular development is able to resume and ovulation is restored [187]. In the ovary, estrogen exerts its mitogenic and differentiative effects through activation of one of two receptors (ER-α or ER-β) belonging to the nuclear receptor family of transcription factors. While G protein-coupled receptor 30 (Gpr30), one of many membrane-bound forms of estrogen receptor, can be detected in the mammalian ovary [239], Gpr30-deficient mice are completely fertile [240]. Following ligand binding in the cytoplasm, ERs dimerize and translocate into the nucleus to activate gene transcription upon recognition of an estrogenresponsive element (ERE) [241]. Esr1 (ER-α) is expressed throughout the female reproductive tract, including in the thecal and interstitial cells of the ovary [242]. The Esr1-knockout mouse is infertile due to reproductive tract defects, including thin uteri, lack of preovulatory follicles, and hemorrhagic cysts in the ovary [242,243]. In contrast,

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ER-β (ESR2) is localized almost exclusively to granulosa cells, and its expression is dynamically regulated by gonadotropins [41,244]. Thus, models of Esr2 disruption have allowed for further understanding of the functions of estrogen in follicular development. ER-β-knockout mice are subfertile with ovaries showing a block in progression from early antral follicles to the large antral and preovulatory stages [245,246]. Granulosa cells of ER-β-knockout follicles show attenuated responses to the differentiating effect of FSH, including suboptimal induction of Cyp19a1—therefore reduced steroidogenesis—and Lhcgr expression [246]. As a consequent of decreased expression of Lhcgr, ER-β-knockout follicles are less responsive to ovulatory LH signals as compared to wild-type controls [247]. These findings suggest that ER-β signaling is important to promote FSHregulated granulosa cell differentiation and promotion of the preovulatory phenotype. Interestingly, ovarian phenotypes of double-knockout Esr1 and Esr2 mice (αβERKO) are completely distinct from either single knockout. αβERKO females are infertile with defective ovarian follicles that appear to transdifferentiate into seminiferous tubule-like structures in adulthood [248]. Physiological and Supraphysiological Effects of Androgens on Follicular Development While initially thought to function solely as the substrate for estrogen synthesis in the ovary [236], the importance of androgens and androgen receptor (AR) signaling in follicular development has now been demonstrated through the generation of granulosa cell-specific knockouts of the AR in mice (ARcKO) [249,250]. Two independently generated ARcKO lines are subfertile, with decreased follicular growth and increased follicular atresia observed starting at the preantral stage [249,250]. Testosterone and dihydrotestosterone (DHT) were shown to prevent granulosa cell apoptosis and follicular atresia by increasing the expression of miR-125b, a known suppressor of proapoptotic proteins [251]. Also in FSH-responsive follicles, androgen increases the level of granulosa cell Fshr mRNA, thus enhancing the FSH stimulation of follicle growth [251]. Testosterone or DHT was also shown to augment FSH-stimulated granulosa cell expression of Star [252]. Clinically, ovarian hyperandrogenism is a feature of the relatively common endocrine and metabolic disorder polycystic ovarian syndrome (PCOS), which often leads to anovulation [253,254]. At supraphysiological levels, DHT treatment of prepubertal rats leads to a block in follicular development at the early antral stage [255]. The abnormally large population of small growing follicles, where dominant follicle(s) fail to emerge, resemble that of PCOS ovaries where these small follicles localize to the ovarian cortex and resemble cysts in diagnostic ultrasound examination. High levels of DHT were shown to inhibit FSH-induced granulosa cell proliferation through

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dampening of PI3K/AKT pathway activation and Cyclin D1 expression [255]. Excess DHT was also shown to inhibit proliferation by preventing insulin- and FSHstimulated activation of the MAPK/ERK and Cyclin D2 expression in cultured granulosa cells [256,257]. These findings demonstrate that at both physiological and supraphysiological levels, androgens influence antral follicular development by modulating both the proliferative and differentiating effects of FSHR signaling.

Conclusions It is abundantly clear that intraovarian signaling pathways are critical for regulating follicular function during both the gonadotropin-independent and -dependent stages of follicle development. Through this review, we hope to have communicated the diversity by which cells of the follicle communicate with each other to ensure coordinated development. It is perhaps not surprising that reproduction, a necessarily robust process to ensure the continuation of species, requires highly interacting, and often overlapping, complex signaling mechanisms. Disruption of early developmental events that are gonadotropin-autonomous, such as follicle formation or control of primordial follicular activation, can directly determine fertility status. However, FSH and LH alone are also not sufficient to support fertility, as their actions are highly dependent upon interactions with underlying paracrine factors. It is remarkable how diverse these intrinsic ovarian regulators are, spanning the gamut of evolutionarily conserved developmental pathways such as Notch and Hedgehog signaling, the large TGF-β family of ligands, many RTKs, and nuclear receptor signaling. The advent of transgenic overexpression and reporter mice, as well as conditional deletion strategies using the Cre/LoxP system in the mouse, has allowed for the discovery of molecular mechanisms that contribute to specific periods of follicular development. In the coming years, we expect that the maturation of CRISPR/Cas9 genome editing technology will extend our knowledge of molecular endocrinology, and perhaps allow relevant questions to be asked in mammals beyond rodents [258]. Moreover, CRISPR/Cas9 technology promises to increase the efficiency—with respect to timing and precision—of generation of transgenic and knockout models. In combination with predictive in silico analysis and high-throughput-omics studies, we may soon identify molecular regulators of poorly understood aspects of follicular development, such as follicle formation and primordial follicle activation. Most findings presented in this chapter represent basic research conducted largely in murine models. We posit that understanding the molecular basis of how ovarian

follicles develop will continue to allow for the identification of the genetic and molecular basis of reproductive disorders, including those of often unknown etiology. Where relevant, we present examples of clinical findings that were informed by these basic studies. The increasing sensitivity and affordability of genome-wide analysis has made possible the identification of genetic polymorphisms in complex, often poorly understood diseases, including idiopathic cases of infertility. Researchers and clinicians continue to identify novel infertility-associated loci, making possible mechanistic studies that are often informed by an understanding of the diverse signaling pathways discussed here. It is an exciting era of discovery, with robust opportunities to continue to increase our understanding of the ovary and its central role in female reproduction.

SUPPORTING GRANTS NIH/NICDH P01 HD021921 NIH/NIGMS T32 GM008061

DISCLOSURE SUMMARY The authors have nothing to disclose

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binding protein 1 (YB-1) enhances gene expression in granulosa cells in response to follicle-stimulating hormone (FSH). J Biol Chem 2016;291(23):12145–60. Cottom J, Salvador LM, Maizels ET, Reierstad S, Park Y, Carr DW, et al. Follicle-stimulating hormone activates extracellular signalregulated kinase but not extracellular signal-regulated kinase kinase through a 100-kDa phosphotyrosine phosphatase. J Biol Chem 2003;278(9):7167–79. Fujinaga H, Yamoto M, Shikone T, Nakano R. FSH and LH up-regulate epidermal growth factor receptors in rat granulosa cells. J Endocrinol 1994;140(2):171–7. El-Hayek S, Demeestere I, Clarke HJ. Follicle-stimulating hormone regulates expression and activity of epidermal growth factor receptor in the murine ovarian follicle. Proc Natl Acad Sci U S A 2014;111(47):16778–83. Babu PS, Krishnamurthy H, Chedrese PJ, Sairam MR. Activation of extracellular-regulated kinase pathways in ovarian granulosa cells by the novel growth factor type 1 follicle-stimulating hormone receptor. Role in hormone signaling and cell proliferation. J Biol Chem 2000;275(36):27615–26. Prasasya RD, Mayo KE. Notch signaling regulates differentiation and steroidogenesis in female mouse ovarian granulosa cells. Endocrinology 2018;159(1):184–98. Wang J, Liu S, Peng L, Dong Q, Bao R, Lv Q, et al. Notch signaling pathway regulates progesterone secretion in murine luteal cells. Reprod Sci 2015;22(10):1243–51. Zhou J, Kumar TR, Matzuk MM, Bondy C. Insulin-like growth factor I regulates gonadotropin responsiveness in the murine ovary. Mol Endocrinol 1997;11(13):1924–33. Zhou P, Baumgarten SC, Wu Y, Bennett J, Winston N, HirshfeldCytron J, et al. IGF-I signaling is essential for FSH stimulation of AKT and steroidogenic genes in granulosa cells. Mol Endocrinol 2013;27(3):511–23. Baumgarten SC, Armouti M, Ko C, Stocco C. IGF1R expression in ovarian granulosa cells is essential for steroidogenesis, follicle survival, and fertility in female mice. Endocrinology 2017;158 (7):2309–18. Park Y, Maizels ET, Feiger ZJ, Alam H, Peters CA, Woodruff TK, et al. Induction of cyclin D2 in rat granulosa cells requires FSHdependent relief from FOXO1 repression coupled with positive signals from Smad. J Biol Chem 2005;280(10):9135–48. Hunzicker-Dunn ME, Lopez-Biladeau B, Law NC, Fiedler SE, Carr DW, Maizels ET. PKA and GAB2 play central roles in the FSH signaling pathway to PI3K and AKT in ovarian granulosa cells. Proc Natl Acad Sci U S A 2012;109(44):E2979–88. Herndon MK, Law NC, Donaubauer EM, Kyriss B, HunzickerDunn M. Forkhead box O member FOXO1 regulates the majority of follicle-stimulating hormone responsive genes in ovarian granulosa cells. Mol Cell Endocrinol 2016;434:116–26. Liu Z, Rudd MD, Hernandez-Gonzalez I, Gonzalez-Robayna I, Fan H-Y, Zeleznik AJ, et al. FSH and FOXO1 regulate genes in the sterol/steroid and lipid biosynthetic pathways in granulosa cells. Mol Endocrinol 2009;23(5):649–61. Miller WL, Auchus RJ. The molecular biology, biochemistry, and physiology of human steroidogenesis and its disorders. Endocr Rev 2011;32(1):81–151. Hillier SG, Whitelaw PF, Smyth CD. Follicular oestrogen synthesis: the “two-cell, two-gonadotrophin” model revisited. Mol Cell Endocrinol 1994;100(1–2):51–4. Conneely OM, Mulac-Jericevic B, Lydon JP, De Mayo FJ. Reproductive functions of the progesterone receptor isoforms: lessons from knock-out mice. Mol Cell Endocrinol 2001;179 (1–2):97–103. Knecht M, Darbon JM, Ranta T, Baukal AJ, Catt KJ. Estrogens enhance the adenosine 30 ,50 -monophosphate-mediated induction

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of follicle-stimulating hormone and luteinizing hormone receptors in rat granulosa cells. Endocrinology 1984;115(1):41–9. Wang C, Prossnitz ER, Roy SK. Expression of G protein-coupled receptor 30 in the hamster ovary: differential regulation by gonadotropins and steroid hormones. Endocrinology 2007;148 (10):4853–64. Otto C, Fuchs I, Kauselmann G, Kern H, Zevnik B, Andreasen P, et al. GPR30 does not mediate estrogenic responses in reproductive organs in mice. Biol Reprod 2009;80(1):34–41. Mangelsdorf DJ, Thummel C, Beato M, Herrlich P, Sch€ utz G, Umesono K, et al. The nuclear receptor superfamily: the second decade. Cell 1995;83(6):835–9. Lubahn DB, Moyer JS, Golding TS, Couse JF, Korach KS, Smithies O. Alteration of reproductive function but not prenatal sexual development after insertional disruption of the mouse estrogen receptor gene. Proc Natl Acad Sci U S A 1993;90 (23):11162–6. Schomberg DW, Couse JF, Mukherjee A, Lubahn DB, Sar M, Mayo KE, et al. Targeted disruption of the estrogen receptor-alpha gene in female mice: characterization of ovarian responses and phenotype in the adult. Endocrinology 1999;140(6):2733–44. Britt KL, Findlay JK. Regulation of the phenotype of ovarian somatic cells by estrogen. Mol Cell Endocrinol 2003;202(1–2):11–7. Krege JH, Hodgin JB, Couse JF, Enmark E, Warner M, Mahler JF, et al. Generation and reproductive phenotypes of mice lacking estrogen receptor beta. Proc Natl Acad Sci U S A 1998;95 (26):15677–82. Couse JF, Yates MM, Deroo BJ, Korach KS. Estrogen receptor-beta is critical to granulosa cell differentiation and the ovulatory response to gonadotropins. Endocrinology 2005;146(8):3247–62. Deroo BJ, Rodriguez KF, Couse JF, Hamilton KJ, Collins JB, Grissom SF, et al. Estrogen receptor beta is required for optimal cAMP production in mouse granulosa cells. Mol Endocrinol 2009;23(7):955–65. Couse JF, Hewitt SC, Bunch DO, Sar M, Walker VR, Davis BJ, et al. Postnatal sex reversal of the ovaries in mice lacking estrogen receptors alpha and beta. Science 1999;286(5448):2328–31.

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[249] Sen A, Hammes SR. Granulosa cell-specific androgen receptors are critical regulators of ovarian development and function. Mol Endocrinol 2010;24(7):1393–403. [250] Walters KA, Middleton LJ, Joseph SR, Hazra R, Jimenez M, Simanainen U, et al. Targeted loss of androgen receptor signaling in murine granulosa cells of preantral and antral follicles causes female subfertility. Biol Reprod 2012;87(6):151. [251] Sen A, Prizant H, Light A, Biswas A, Hayes E, Lee H-J, et al. Androgens regulate ovarian follicular development by increasing follicle stimulating hormone receptor and microRNA-125b expression. Proc Natl Acad Sci U S A 2014;111(8):3008–13. [252] Laird M, Thomson K, Fenwick M, Mora J, Franks S, Hardy K. Androgen stimulates growth of mouse preantral follicles in vitro: interaction with follicle-stimulating hormone and with growth factors of the TGFβ superfamily. Endocrinology 2017;158(4):920–35. [253] Franks S. Polycystic ovary syndrome. N Engl J Med 1995;333 (13):853–61. [254] Ehrmann DA. Polycystic ovary syndrome. N Engl J Med 2005;352 (12):1223–36. [255] Chen M-J, Chou C-H, Chen S-U, Yang W-S, Yang Y-S, Ho H-N. The effect of androgens on ovarian follicle maturation: dihydrotestosterone suppress FSH-stimulated granulosa cell proliferation by upregulating PPARγ-dependent PTEN expression. Sci Rep 2015;5(1):18319. [256] Kayampilly PP, Menon KMJ. Dihydrotestosterone inhibits insulin-stimulated cyclin D2 messenger ribonucleic acid expression in rat ovarian granulosa cells by reducing the phosphorylation of insulin receptor substrate-1. Endocrinology 2006;147 (1):464–71. [257] Kayampilly PP, Menon KMJ. AMPK activation by dihydrotestosterone reduces FSH-stimulated cell proliferation in rat granulosa cells by inhibiting ERK signaling pathway. Endocrinology 2012;153(6):2831–8. [258] Wang Y, Du Y, Shen B, Zhou X, Li J, Liu Y, et al. Efficient generation of gene-modified pigs via injection of zygote with Cas9/ sgRNA. Sci Rep 2015;5(1):8256.

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C H A P T E R

3 Human Folliculogenesis Revisited: The Menstrual Cycle Visualized by Ultrasonography Roger A. Pierson INTRODUCTION

noninvasive visualization of the inner workings of the reproductive system possible. Our enhanced understanding of ovarian function during the reproductive cycle is now synergistic using data from our knowledge of microstructure, elaboration of the reproductively active hormones, selection of optimal animal models, and the images generated by ultrasonography. It is by blending these direct and indirect sources of information that we are able to build new conceptual models to understand ovarian function from puberty, through the reproductive years to reproductive senescence. The development of transabdominal ultrasonography in the late-1970s granted the ability to assess the serial growth and regression of follicles [19–27]. The advent of high-resolution transvaginal ultrasonography in the late 1980s dramatically improved our ability to visualize ovarian structures in situ. Antral follicles as small as 1 mm can now be detected and the growth dynamics of individually identified follicles described (Fig. 1) [28–35]. Ultrasonography allows us the ability to visualize ovarian structures and the changes that they undergo over time. The objective of this chapter is to provide a view of human ovarian folliculogenesis generated by the synergy of imaging the ovaries, endocrinologic assessments, and the insights gained from experiments in animal models.

To visualize means “to see.” In reproductive medicine, we are able to use constantly evolving visualization technologies to elucidate the anatomic processes that underly reproduction. For millennia, we have looked at the outside of the body and wondered what was going on inside. The role of the ovaries as the producer of eggs has been actively explored since the time of William Harvey who proclaimed “ex ovo omnia” (all things originate from eggs) through the Long 18th century [1]. The drawings and insights of Andreas Vesalius, Girolamo Fabrici, William Harvey, and Regnier de Graaf represented the standard of knowledge. However, these pioneers were limited by the lack of the best tools to elucidate the detailed structure of the ovary. It was not until 1837 that Karl Ernst von Baer discovered that the mammalian oocyte was enclosed within the ovarian follicle [1]. It was then not until the 20th century that the first descriptions of human ovarian follicular development were made [2,3]. It has been the technologic advances in imaging over the past 60 years that have provided noninvasive tools for evaluating ovarian function in women. Before were able to use direct visualization of the female reproductive organs, the conceptual models for understanding ovarian function over the menstrual cycle were developed from observations obtained using a combination of visual anatomical techniques and inference from endocrinologic techniques. Many of our notions of how the ovaries work over time have been extrapolated from anatomic studies in nonhuman primates, domestic animals, and laboratory animal species. Early studies to elucidate human ovarian and menstrual cyclicity in the 1950s–70s were based on histologic and/or endocrinologic assessments [2,4–18]. Only recently has the technology existed to make

The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00003-0

FOLLICULOGENESIS The Language of Folliculogenesis The processes of folliculogenesis in the medical/ scientific literature are described in differing terminology in different studies and among different species. For example, in the human literature, the term “recruitment” has

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© 2019 Elsevier Inc. All rights reserved.

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FIG. 1 Normal ovaries (A, B) during a natural menstrual cycle demonstrating normal follicle population and distribution on day 12 post menstruation. A dominant follicle is visualized in the central portion of the image (A) and several subordinate follicles from the wave (2–5 mm) are observed in the left lateral aspect of the ovary. A normal population of 4–5 mm follicles is observed in the contralateral ovary (B). Gently scanning the ovaries from medial to lateral aspects allows quantitation of the follicle population and facilitates ovarian mapping. Daily scanning and mapping allows determination of the fates of individual follicles.

been used to describe three distinctly different physiologic events: (1) the initial transition of primordial follicles from the resting pool into the preantral growth phase, (2) the cyclic recruitment of a cohort of small antral follicles (2–5 mm) during the menstrual cycle following puberty, and (3) the preferential growth of the dominant ovulatory follicle. Follicles comprising a recruited cohort of 2–5 mm follicles have been referred to as “selectable” follicles [36]. Similarly, the term “selection” has been used to describe two different phenomena: (1) the recruitment of a cohort of small antral follicles and (2) the preferential growth of a species-specific number of large antral follicles from the recruited cohort. The follicle that is selected from the recruited cohort has been referred to as “dominant” [37] or “privileged” [38], while other follicles of the cohort that undergo atresia have been termed “ordinary” [38], “challenger” [39], “subdominant” [40], or “subordinate” [41]. For the purposes of this chapter, it is important to maintain consistency with the terminologies used in the human and animal literature, other chapters in the present volume, and a recent review of human folliculogenesis [35]. Follicle “recruitment” and “selection” have been used to represent two different physiologic events. “Recruitment” refers to the emergence of a group or cohort of small (2–5 mm) antral follicles while “selection” refers to the preferential growth of a dominant follicle from the cohort of recruited antral follicles. “Subordinate” follicles comprise all follicles of the recruited cohort, excluding the dominant follicle. “Cohort” is used interchangeably

with “wave.” The terms “recruitment” and “emergence” are used interchangeably. The term “interovulatory interval” (IOI) has emerged from the language of detailed studies of folliculogenesis in domestic animals and is used to define the time period between successive ovulations [42–45]. Ovulation represents the terminal event in the life of a follicle that has been privileged to express both its endocrine and its exocrine function. This term has also been used in studies designed to elucidate human ovarian function [35,42,46,47]. It makes sense to describe the human ovarian cycle from ovulation to ovulation to be consistent in discussions of critical endpoints and concepts between human and animal model studies. Menses can be regarded as an endometrial reflection of ovarian function in the event that conception does not occur. It is important to note that the ovarian cycle described by the IOI and the menstrual cycle describing the events between menses are chronologically offset by approximately 14 days.

Preantral and Early Antral Follicle Development Preantral follicles and the earliest stages of antral follicle development are beyond the resolving power of the current state-of-the-art ultrasound instruments. We will not be able to image these very early stages of folliculogenesis over time until a means of noninvasive “virtual histology” is developed. Briefly, human folliculogenesis, from the primordial phase to the preovulatory phase, has

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been estimated to span approximately 175 days [38]. Follicular development in the ovaries of female fetuses begins as early as the fourth month post conception [9]. The primordial germ cells have migrated from the yolk sac endoderm to the gonadal ridge and increase in number by mitotic division. The first meiotic division occurs when the germ cells arrive at the gonadal ridge—the oogonia enter the tissues comprising the ridge, become primary oocytes, and in the process form the primitive gonad. Somatic cells (i.e., surface epithelial cells, follicular granulosa and theca cells, interstitial cells, fibroblasts) surround the oogonia and form rudimentary ovarian follicles ( 2 mm, onset of ovulation, and menstrual cyclicity (reviewed in Refs. [56,57]).

The Prepubertal and Pubertal Period Human follicular development to the early antral stage occurs during infancy and childhood [58]. Those follicles become atretric and regress; the stimulus for their

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recruitment and development to the antral stage remains unclear. Similarly, the first few menstrual cycles during puberty may be anovulatory due to incomplete maturation of the hypothalamo-hypophyseal axis and the inability of early pubertal pituitaries to produce sufficient LH to trigger the cascade of events leading to ovulation. In animal models, sequential waves of anovulatory antral follicle development have been documented in prepubertal calves, as early as 2 weeks of age [59,60]. The transition to the equivalent of puberty in calves does not appear to have been studied in detail. Our understanding of follicular dynamics in prepubertal and pubertal girls remains poorly elucidated due to ethical and technical constraints on conducting reproductive research imposed by the age and ability to consent of the girls in this population. In many ways, it would be highly beneficial to our understanding of pubertal disorders and the effects of various pharmaceutical interventions and environmental insults to know how the ovary first matures to first ovulation and precisely how subsequent adult ovarian cyclicity is established.

Recruitment of Antral Follicles Our understanding of how and when antral follicles are recruited has been limited by the technologies that we are able to bring to bear on this fundamental issue. Antral follicles 2–5 mm in diameter are detected histologically in excised specimens and ultrasonographically throughout the human menstrual cycle upon clinical investigation [29,46,61,62]. The pattern of emergence of 2–5 mm follicles, however, is a matter of long-standing debate. It is noteworthy that 1–2 mm represents the lower limit of resolution of the current generation of diagnostic ultrasound instruments. Three distinct theories of follicular recruitment have been proposed and have been reviewed in detail in Ref. [35]. They are (1) a Continuous Recruitment Theory and (2) Cyclic Recruitment Theories. The Cyclic Recruitment Theories are subdivided into (1) a single episode of recruitment per ovarian cycle in which a cohort of 2–5 mm follicles is recruited from a continuous supply of antral follicles once during each menstrual cycle [8,23,29,61] and (2) wave patterns of recruitment in which multiple cohorts or “waves” of antral follicle recruitment occur [2,21,24,42,46,63]. In the single episode model, a single increase in the number of follicles 2–5 mm would be detected after regression of the CL in the late luteal phase or early follicular phase of the menstrual cycle. The single episode has been referred to as the “privileged” phase of follicle development. In the wave model, a “wave” of follicular development is defined as the synchronous growth of a group of antral

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follicles that occurs at regular intervals during the menstrual/estrous cycle; follicles in each wave are of similar, but not identical, diameters [43,46,64,65].

Folliculogenesis During the Interovulatory Interval A “wave” of follicular development is defined as the synchronous growth of a group of antral follicles that occurs at regular intervals during the IOI or menstrual/ estrous cycle; follicles in each wave are of similar, but not identical, diameters [35,43,46,64,65]. Each follicle wave is composed of a group of antral follicles with synchronous growth [66]. Typically, one follicle grows to a larger diameter and becomes the lead, dominant follicle of the cohort. New follicle waves appear at regular intervals within cycles and each of the waves is preceded by a small increase in FSH. Within each cycle, the earlier waves are consistently anovulatory whereas the final wave ends with ovulation. The earliest studies of human follicular dynamics in which waves of follicle development were inferred involved histologic evaluations of excised ovaries removed at different times of the menstrual cycle [2]. Two waves of follicle growth (follicles > 1 mm) were surmised during the menstrual cycle based on the microstructural status of the follicles. The first wave occurred in the follicular phase and a second wave occurred in the luteal phase. Physiologically healthy antral follicles >10 mm were detected only during the follicular phase in most women, but some were observed during the late luteal phase [4]. Luteal phase follicles had fewer granulosa cells and produced less estradiol compared to those in the follicular phase [4,5]. In another study, more antral follicles >1 mm were detected in the luteal phase [67]. Most follicles were atretic; however, more healthy antral follicles >1 mm were observed in the early luteal vs mid-luteal and late follicular phases. This observation was interpreted to mean that follicles originating from different waves of development then in various stages of atresia remained in the ovaries. In an early ultrasound study of ovarian function performed using transabdominal ultrasonography, antral follicles could be identified in the ovaries, the development of the preovulatory follicle followed and ovulation detected [21,63]. Two periods of antral follicle development were detected in women with regular 30–35 day cycles compared to one follicular wave in women with 26–30 day cycles. Following these reports, most attention was focused on the evolution of the preovulatory follicle and how it varied from atretic follicles from the same cohort [68]. In the most detailed study of its kind, the ovaries of 63 clinically normal women between the ages of 19 and 43 (mean age 28 years and 7 months) were examined

daily using high-resolution transvaginal ultrasonography [66,69]. The objective of the study was to characterize the daily growth and regression of ovarian follicles in women during one IOI to evaluate the wave-like changes in the number and diameter of follicles during an IOI. At each examination, all follicles > 2 mm were counted and measured. The locations of each follicle >8 mm were also sketched on an ovarian map in order to facilitate following the fates of individual follicles. The day-to-day identities of individual follicles were determined using the internal iliac blood vessels, the ovarian hilus, and the location of neighboring follicles and the corpus luteum within the ovary as landmarks. The diameter profiles of individual follicles that grew to > 8 mm throughout the IOI were graphed for each woman. Ovulation was defined as the disappearance of a large follicle (> 15 mm) that had been identified by ultrasonography on the previous day and the simultaneous visualization of a luteal structure [70]. Wave emergence was defined as the day on which the largest follicle of the wave was detected at 4–5 mm. Five distinct patterns of follicular development were observed [66,69]. Emergence of waves of between 4 and 14 follicles > 4–5 mm were detected either two or three times during the IOI. Most women (68%) exhibited distinct two waves of follicle recruitment during the IOI, while the remaining women (32%) exhibited three waves. The mean IOI of women with three follicular waves was 29 days— significantly longer than the mean IOI of 27 days in women with two waves. Wave patterns were subdivided into major waves and minor waves. Major waves were defined as those in which a dominant follicle was selected and experienced preferential growth. Minor waves were those in which divergence and dominance were not observed. Most women developed a major ovulatory wave in the follicular phase and one or two minor anovulatory waves in the preceding luteal phase. The final wave resulted in ovulation of the dominant follicle of a major wave [66,69]. Women with two follicular waves exhibited an anovulatory wave of follicles that emerged at the time of ovulation (i.e., early luteal phase). The luteal phase wave was followed by emergence of the ovulatory wave during the early follicular phase; that is, at the time. In women with three wave patterns of folliculogenesis, an anovulatory wave emerged at the time of ovulation, a second anovulatory wave emerged during the mid- to late luteal phase, and a third ovulatory wave emerged in the early-mid follicular phase [66,69]. Emergence of the ovulatory wave consistently followed a decrease in diameter of the dominant follicle(s) of the preceding anovulatory wave. Observations of follicular waves in women are consistent with those previously documented in several animal species, including cattle [43,65,71], mares [64,72–75], sheep and goats [76,77], llamas and alpacas [78,79], musk oxen [79a], water buffalo [80], as well as deer and wapiti

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[81,82]. The day of follicular wave emergence is recognized ultrasonographically when the largest follicles of the cohort first attain diameters of 4–6 mm in women [46,64,66,69], compared to 4–5 mm in cows [71,83,84], 4–5 mm in sheep [77], and 13–15 mm in mares [64,73]. Evidence of follicular waves has also been reported in subhuman primates [85,86]. The emergence of both major and minor follicular waves was preceded by an increase in FSH that occurred in concert with an increase in the number of follicles >5 mm. The nadir in FSH, which preceded wave emergence, occurred 2 days earlier in women in which major vs minor follicular waves were observed. Estradiol levels increased earlier during the follicular phase in women with two follicular waves. This observation was attributed to earlier emergence of the dominant follicle. Similarly, preovulatory surges of FSH, LH, and estradiol occurred earlier in women with two vs three follicle waves: women with two waves had shorter IOI. Differences were not observed in the concentrations of estradiol and LH in women exhibiting major vs minor patterns of follicle-wave development. Nor were differences observed relative to the location of the corpus luteum. In all of the women studied, the final follicular wave of the IOI was an ovulatory major wave; preceding waves were either minor or major anovulatory waves. The diameter of the dominant preovulatory follicle was larger than that observed in dominant follicles of anovulatory major waves. Selection of the dominant follicle in both major anovulatory and ovulatory waves occurred at a diameter of approximately 10 mm, which was consistently 3 days after follicle wave emergence [66,69]. The natural history of wave patterns of folliculogenesis during the cycle provides an explanation for persistence of large anovulatory follicles in the early follicle phase. A large anovulatory follicle may begin growth within a luteal phase wave and persist as the next wave of follicles develops. Serial imaging in the early to mid- or latefollicle phase can be used to map the fate of the follicle as it grows, becomes atretic, and regresses spontaneously with time. Understanding that folliculogenesis occurs in regular wave patterns also provides an explanation for some women to have consistently longer menstrual cycle lengths, as women with >2 follicle waves had progressively longer cycles than women with only two wave cycles. Variability in cycle lengths and the formation of dominant anovulatory follicles in the luteal phases of normal menstrual cycles can now be recognized as physiologically normal processes.

Dominant Follicle Selection “Selection” is the process by which a single “dominant” follicle is chosen from the recruited cohort or wave for preferential growth [2,29,37,38,46,61,87–89]. Selection has been described as a phenomenon of

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avoiding atresia. That is, the selection process may be considered a hierarchical progression of follicle atresia over the period encompassing the rise and fall in FSH. The smallest follicles in the cohort are the least capable of competing for diminishing levels of FSH and therefore undergo atresia first, followed by regression of progressively larger follicles in the wave until ultimately only one (dominant) follicle survives [90,91]. Selection of a dominant follicles has been generally described to occur once in the early to mid-follicular phase of the menstrual cycle and it is expected that the selected follicle becomes dominant and ends its life as a follicle at ovulation [29,62,92,93]. However, selection of dominant follicles has now been observed to occur two or three times during natural menstrual cycles in healthy women [35,66,69]. Dominant follicles were observed to develop and subsequently regress prior to selection of the ovulatory follicle in nearly a quarter of natural menstrual cycles. Reports of anovulatory dominant follicles developing spontaneously prior to the ovulatory follicular wave in some, but not all, women are strikingly similar to follicular dynamics in mares [64], despite differences in luteal phase lengths between species.

Follicle Divergence The dominant and largest subordinate follicles of the ovulatory wave undergo a common growth phase in women, consistent with observations in mares and cows [83,87,94]. Selection is manifest when the growth profile of the dominant follicle begins to “diverge” as it continues to grow while subordinate follicles undergo atresia [87,90]. In women, divergence occurs when the dominant follicle reaches a diameter of
10 mm on day 6–9 of the follicular phase [29,46,87,88]. It is important to consider that the physiologic events that underlie divergence may well occur minutes or hours before the actual observation of changes in the diameter of the dominant follicle relative to others in the cohort. The changes in the growth dynamics of dominant and first subordinate follicles in women are similar to that described in other monovular species; however, the detailed investigation of periselection folliculogenesis has not been performed [87,95].

Unified Theory of Selection A recently proposed theory of selection posits that the largest follicle in the wave does not exert dominance over other follicles during the common growth phase before deviation is initiated. The models for selection are most fully developed in mares and cattle [83,94]. Deviation, as observed by differential changes in the growth rates of future dominant and subordinate follicles, begins approximately 24 h prior as evidenced by structural changes in the follicle walls and increased vascular flow to the future

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dominant follicle. The model proposes that the deviation process represents the complete mechanism of dominant follicle selection. The observations of selection and deviation in women are conceptually similar; however, detailed studies to confirm the mechanism have not been performed. The follicle diameters at which deviation is manifest in women is between approximately 8.5 and 10 mm [35]. If the unified theory is correct, follicle deviation and follicle selection represent the same physiologic process. This information may represent a significant advance in the design of new ovarian stimulation protocols.

Follicle Dominance Conceptually, to be dominant, a follicle must suppress the growth of subordinates of the same wave and suppress emergence of the next follicular wave through an intraovarian or systemic inhibitory effect [90,96]. Observations from human and animal studies support the idea that the dominant follicle exerts both morphologic and functional dominance once selection has occurred. Selection is manifest in preferential development of the dominant follicle following deviation. It has been postulated that the future dominant follicle may contain more granulosa cells and FSH receptors, making it more sensitive to FSH, compared to the subordinate follicles [97]. Subordinate follicles are not able to thrive in an environment of declining FSH and therefore succumb to atresia [46,88,98–101]. The functional status of dominant follicles that develop during anovulatory waves preceding the ovulatory wave in women is not fully understood. However, the insights provided by the equine model would seem an appropriate starting place, given the similarities between mares and women in the processes of folliculogenesis [64,75,89]. Estradiol levels have been shown to increase in association with the emergence of anovulatory follicle waves during the luteal and early follicular phases of the cycle [46]; however, the source of the estradiol (follicle vs corpus luteum) is not known. In a preliminary study, no differences in circulating estradiol concentrations were observed in the luteal phases of women with anovulatory major waves vs minor waves [75]; however, data from a larger sample of women are required to either confirm or refute the preliminary inference. Ultrasonographic image attributes of the dominant follicles have been found to differ between anovulatory and ovulatory waves [102]. Taken together, these data suggest that dominant follicles of anovulatory waves may exhibit different physiologic characteristics than dominant follicles of ovulatory waves in women. The morphologic and endocrinologic changes occurring in association with the development of follicle waves during the human menstrual cycle are illustrated (Fig. 2).

Luteal Influences on Follicle Development and Selection The idea that the corpus luteum inhibits the development of antral follicles is challenged by the documentation of multiple follicle waves during the menstrual cycle and the potential for development of more than one dominant follicle. A deeper investigation is required into the specific role of the CL on follicular dynamics. For example, the evidence of aromatase activity in the CL of mares that provides rationale for the hypothesis that the CL of domestic animals also produce estradiol and its possible roles in folliculogenesis at different times during the IOI [103]. It is also important to elucidate the role of the mid-luteal increase in estradiol that occurs during anovulatory waves: it is not known if the estradiol in women is of follicular or luteal origin. Similarly, the role of luteal inhibin in regulating antral folliculogenesis in women remains unclear. The role of the CL in regulating follicular wave dynamics has been studied in women and domestic farm animals. No differences were observed in the size or life span of the CL, progesterone secretion, or luteal phase estradiol secretion in women who exhibited two vs three waves or in women who exhibited major waves vs minor waves of follicular development prior to the ovulatory wave [104]. However, the CL appeared to influence dominant follicle selection in women with three follicular waves. When the second of three waves emerged in the mid-luteal phase, selection of a dominant follicle did not occur; that is, a minor wave developed [46]. In comparison, when the second wave emerged in the late luteal or early follicular phase, a dominant follicle was selected; that is, a major wave developed [46,66,69]. The role of the CL in follicle wave emergence and selection is the subject of intense scrutiny in domestic animal models [64,75,84,89,105–108]. The choice of optimal experimental model to elucidate these challenging concepts is yet to be made selected; however, primates, including humans, are likely best modeled by mares and cattle. In humans, studies to determine the role of the CL and its influence on dominant follicle selection are limited. It was observed that seven out of eight dominant follicles developed contralateral to the CL from the previous ovulation in women [61,66,69]. Dominant follicles that developed contralateral to the previous ovulation exhibited higher estradiol/androstenedione ratios than dominant follicles that developed ipsilateral to the previous CL [109]. In a clinical study, pregnancy rates following insemination and IVF in natural cycles were found to be higher when the ovulatory follicle developed contralateral to the CL from the previous ovulation [109]. The limitations of the observational study design and small sample sizes make interpretation of these findings difficult. However, evaluations of ovulation in both infertile

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FIG. 2 Schematic representation of changes in the morphologic and endocrinologic dynamics associated with 2 (A–C) and 3 (D–F) waves of follicle development in women. Data are presented over 1 complete interovulatory interval (IOI) and 1 complete menstrual cycle for illustrative purposes. Dotted vertical lines indicate the days of wave emergence (W1 ¼ Wave 1, W2 ¼ Wave 2, W3 ¼ Wave 3). Follicles are shown in pink and the CL is shown in yellow (A, D). Major anovulatory waves (ghosted) were detected in some, but not all, women prior to the ovulatory wave (A, D). Changes in serum concentrations of FSH and LH (B, E) and estradiol and progesterone (Continued)

and fertile women support the notion that follicle selection and subsequent ovulation of the dominant follicle occur randomly between the right and left ovaries as observed in domestic animal species [46,110–114].

Preovulatory Follicle Development The dominant follicle of the final follicular phase wave develops preferentially after selection and it typically reaches preovulatory status at a diameter of 16–29 mm in the late-follicular phase [20,25,27,29,33,38,42]. The preovulatory follicle grows at a rate of 1–4 mm/day, with reports of increases, decreases, or no change in growth

rate in the few days leading up to ovulation. Preovulatory follicles grow slightly faster after selection than before [20,21,29,33,68,115–118]. Preferential growth of the dominant follicle in the midto late-follicular phase is associated with increased aromatase activity and a rapid elevation of circulating and follicular fluid estradiol-17β [5,21,25,33,46,61,119,120]. Increased gonadotropin responsiveness in the dominant follicle is presumed to be responsible for mediating dominant follicle granulosa cell estradiol production, LH receptor expression, and continued preovulatory growth [88,120–126]. The dominant follicle is responsible for over 90% of the estrogen production in the preovulatory period [13].

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FIG. 2 CON’D (C, F) are to facilitate elucidation of the chronology of the reproductive cycle. (Reprinted with permission from Baerwald AR, Adams GP, Pierson RA. Ovarian antral folliculogenesis during the human menstrual cycle: a review. Hum Reprod Update 2012;18(1):73–91.)

Both intraovarian and endocrine factors contribute to and regulate preovulatory follicle growth. These factors and their role in folliculogenesis are discussed in detail in other chapters in this volume. Briefly, the discussion centers around (1) increased granulosa cell aromatase mRNA expression, (2) follicular fluid anti-Mullerian hormone (AMH) concentrations, (3) stimulation of inhibin A from thecal cells increasing androgen production, (4) IGF-II mRNA expressed by granulosa cells, and (5) stimulation of aromatase activity [99,101,127–130]. The oocyte and follicle are in constant communication and oocytederived factors GDF-9 and BMP-15 appear to be required for follicle development to the ovulatory stage and MP-15 may be involved in expansion of the cumulus oophorous [131,132]. Estradiol production from the dominant follicle peaks the day before the LH surge providing positive feedback at the hypothalamus and pituitary stimulating the surge of LH required for inducing ovulation [119,133]. The

preovulatory follicle is highly vascularized, has acquired LH receptors, and is then able to respond to the mid-cycle rise in LH [4,119,134]. Ovulation occurs, on average, within 24 h of the LH peak [25]. Systemic progesterone concentrations begin to rise after the preovulatory estradiol peak but before the LH surge and indicate the onset of luteinization [4,61,119,135,136].

Follicle Selection in Anovulatory Waves Selection of a dominant follicle was once thought to occur only once during the menstrual cycle [29,62,93]. That is, at the time of physiologic selection of the preovulatory follicle. However, selection of a dominant, anovulatory follicle occurs more than once in approximately one-quarter of apparently healthy women [35,66,69]. The findings in humans are similar to domestic farm animals, particularly, mares [64,72]. Ovarian follicular

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dynamics in women of reproductive age is clearly more variable than previously thought. The direct observations on ovarian function facilitated by ultrasonography require detailed investigation of the functional status, oocyte quality, and ovulatory potential of follicles that comprise anovulatory follicular waves. Folliculogenesis and ovulation in mammals also are influenced by environmental factors such as energy balance, body condition, photoperiod, temperature, and exposure to endocrinedisrupting chemicals; however, the effects of environmental factors on ovarian function are poorly elucidated [137–141].

Repeatability in Follicular Wave Patterns The consistency, or lack thereof, in the number of waves that develop per cycle has clinical implications for optimizing strategies that manipulate ovarian follicular development. The need for knowledge is driven by desire for the development of technologies as varied as safer, more effective hormonal contraception and ovarian stimulation regimens that yield optimal numbers of fertilizable oocytes. Similarly, knowledge of how wave patterns of follicular develop during puberty is crucial in understanding normal folliculogenesis in women during their reproductive years, and who have polycystic ovary syndrome or other metabolic conditions. Elucidation of patterns of folliculogenesis as women transition from their reproductive years to reproductive senescence is clearly important in understanding conception and contraception needs as women age. At present, the number of follicle waves per estrous cycle in cattle is consistent within individuals [142,143]. Similar work in humans has not been reported. Research on the repeatability of wave patterns within individual women over time in both long-term and short-term periods is logistically challenging; however, this knowledge is fundamental for designing studies to elucidate the mechanisms underlying follicular wave dynamics.

Transition to Reproductive Senescence Age-related changes in antral follicle dynamics remain poorly elucidated. Research to characterize age-related changes in follicular wave dynamics may help to explain the earlier selection of the dominant follicle and shorter follicular phases, as well as abnormal folliculogenesis, ovulation, and estradiol secretion observed in women of advanced reproductive age [47,144–146]. Age-related decreases in ovarian reserve correlate with an observed decrease in the number of antral follicles 2–10 mm in diameter [147]. A mechanistic approach to elucidating the observations is that a decrease in the

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AFC in peri-menopausal women results in a reduction in circulating inhibin B, AMH, and IGF-I and a coincident increase in FSH and cycle irregularity. Circulating estradiol levels remain the same or are elevated [147–157]. Continuing depletion of the number of antral follicles in the ovaries results in continued decline in inhibin B, thereby increasing FSH and LH. Estradiol and progesterone decrease and the loss of menstrual cyclicity, at first sporadically, culminating in menopause when women reach approximately 51 years of age (reviewed in Ref. [158]). The direct relationships between age-related changes in antral follicle dynamics and hormone production were explored to determine if age-related changes in antral follicle dynamics would be associated with changes in hormone secretion as women age [47]. Women in their mid-reproductive years (MRA; 18–35 years) and advanced reproductive years (ARA; 45–55 years) were examined three times weekly for a complete IOI. The growth dynamics of follicular phase dominant follicles did not differ in women of mid- vs advanced reproductive age [7]. However, differences in follicular phase hormone patterns were detected in association with luteal phase dominant follicles. Luteal phase dominant follicles (LPDFs) in mid-reproductive age women were associated with increased luteal phase inhibin B and estradiol. Other aspects of the endocrine system were not different among women in the two age groups. Early follicular phase estradiol was greater in older women who developed atypical LPDF compared with those who did not. In reproductive age women, luteal phase dominant follicles were a product of normal follicular activity and were associated with elevations in luteal phase estradiol. Luteal phase dominant follicles in the older group of women emerged earlier, grew larger, persisted longer and were associated with acute and atypically high luteal-follicular phase estradiol. Progesterone concentrations were lower in the older women with luteal phase dominant follicles vs those without dominant follicles in the luteal phase of their cycles. As the interval to menopause decreases, ovarian activity becomes increasingly sporadic, Selection of a dominant follicle appears to occur earlier during the IOI and shorter follicular phases are observed. These changes lead to shorter intermenstrual intervals, anovulation, and an increased incidence of dizygotic twinning in women who conceive [154,159–163]. Earlier selection of a dominant ovulatory follicle in aging women has been attributed to a faster growth rate of the dominant follicle or earlier emergence of the follicular cohort in the luteal phase of the preceding cycle [164,165]. Similar age-related changes in ovarian function have been reported in mares and cows [166–169].

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HORMONAL INFLUENCES ON FOLLICULOGENESIS Detailed discourse of the hormonal influences on, and control of, folliculogenesis is presented elsewhere in this volume. The following is intended as a very brief discussion to relate associations and context among

observations made via focused experimentation to those visualized by serial visual analyses made using ultrasonography (Fig. 3). An elevation in circulating FSH appears to precede the recruitment of each follicular wave during the IOI in women, which is consistent with previous reports in domestic animals [41,46,66,69,76,96,170]. The height of the peak in

FIG. 3 Sequence of ultrasonographic images of ovulation in an individual woman. Follicular evacuation occurred over 11 min from the first detected fluid leakage to complete apposition of the follicle walls. Images range from 1 min prior to the onset of ovulation to complete follicular evacuation and represent the immediately preovulatory follicle (upper left image) and first fluid leakage from the stigma (upper middle image). The remainder of the images represent 10% declinations in fluid volume leading to complete follicular evacuation (lower right image). Time-code values are shown in the lower left corner of each image displaying hours, minutes, seconds, and frame number. (Reprinted with permission from Hanna M, Chizen D, Pierson R. Characteristics of follicular evacuation during human ovulation. Ultrasound Obstet Gynecol 1994;4:488–93.)

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circulating FSH in cows is inversely related to the number of follicles recruited into a wave [143,171]. The inverse relationship between FSH and number of follicles in a wave cannot be directly explained by alterations in the secretion of estradiol, inhibin, or IGF-I; however, it is consistent with the observations of age-related decrease in AFC and elevated serum FSH in women [154,159,171], cattle [169], and sheep [172]. The increase in circulating FSH is a requisite for eliciting recruitment of the follicular cohort or wave [46,88,96,173,174]. It appears to be the duration of the rise in FSH above a critical threshold that determines the number of dominant follicles selected from the recruited cohort for preferential growth [175,176]. It is the nature of the postsurge decline in FSH that is a critical factor in selection of the dominant follicle [90,96]. This concept has been termed the “FSH Threshold/Window/Gate Concept” [110,175,177]. If the duration that FSH is above the threshold is short, a single dominant follicle will develop. However, if FSH is above the threshold for a longer period of time, multiple follicles may be selected at the same time. The association with this notion is surmised from observations in polyovular species and during ovarian stimulation therapies in women [175]. The precise roles of inhibin A, inhibin B, and AMH in regulating the emergence of multiple follicular waves in women remain elucidated. All follicles of the cohort produce inhibin B that contributes to the decrease in FSH that occurs prior to selection [176,178,179]. Inhibin B inhibits continued FSH secretion in the mid-follicular phase [88,180]. A second short-lived peak in inhibin B has been documented 2 days after the mid-cycle LH surge providing support for the observations of luteal phase waves of follicular growth in women [180]. The role of activin in dominant follicle selection in women is not well elucidated. AMH continuously decreases in follicles from 3 to 12 mm, which encompasses the time during which selection occurs [89,94,181]. Aromatase activity begins in granulosa cells of follicles larger than 6–8 mm (typically day 5–8 of the menstrual cycle). The dominant follicle produces more estradiol17β than other follicles in the cohort; while atretic subordinate follicles exhibit a greater androgen/estrogen ratio in their follicular fluid [36,61,120,135,182–187]. LH-induced production of androgens in thecal cells provides the substrate for estradiol production in granulosa cells [110,188,189]. Estradiol-17β from the dominant follicle provides negative feedback on pituitary FSH secretion which contributes, in part, to the mid-follicular phase decrease in circulating FSH and inhibition of subordinate follicle growth [125]. LH receptors form on granulosa cells of the dominant follicle following estradiol secretion [190,191]. The dominant follicle becomes less dependent on FSH and more responsive to LH during selection [122,123,192]. Differences and similarities between the

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secretion of gonadotropins, inhibin, and estradiol during follicle selection in women, compared and contrasted to that in domestic farm animals have been reviewed in detail in Refs. [83,87,89,94,193]. AMH may play a role in determining the number of recruited follicles in a wave. AMH is produced from the granulosa cells of primary, secondary, preantral, and early antral follicles (< 4 mm) and acts to inhibit the initiation of primordial follicle growth from the ovarian reserve [194,195]. Although not fully elucidated, AMH may play a role in regulating recruitment of the antral follicular cohort [149,196]. In mice, AMH decreases the sensitivity of follicles to FSH thereby inhibiting FSH-induced antral follicle growth [197]. The insulin-like growth factor (IGF) system plays an important role in the intraovarian regulation of antral follicular development. Circulating levels of IGF-I and IGF-II do not appear to differ during the human menstrual cycle [198]. However, changes in follicular fluid IGF concentrations have been reported [199]. The role of IGF-I and II in regulating antral follicle development in women appears to be quite similar to that described in rodent and domestic animal species. In addition, there is evidence in rodents to suggest that follicular activin, GDF-9, AMH, and BMPs may regulate dominant follicle selection by modulating granulosa cell IGF-dependent signaling pathways [105,200,201] (reviewed in Ref. [202]).

OVULATION AND LUTEOGENESIS Ovulation, by definition, is the rupture of a preovulatory ovarian follicle, evacuation of the follicular fluid, and expulsion of the oocyte (Fig 3). It is the most dramatic event in the life of a follicle. Transvaginal ultrasonography has made it possible to capture ovulation in real-time [203,204]. The time required for ovulation varied from less than 1 min to more than 20 min from the initial fluid leakage to the complete apposition of the follicle walls. Evacuation of follicular fluid during ovulation averaged approximately 10 min. The site of ovulation could be identified as soon as ovulation occurred by examining the external surface of the ovary for the point of rupture. After ovulation, the cells of the former follicle must reorganize, both structurally and functionally, to form the luteal gland. Luteogenesis involves invasion of the former follicle walls by newly developing vascular channels and the granulosa cells, thecal cells, and invading vasculature combine to become the new corpus luteum. The CL will produce the progesterone necessary to maintain pregnancy should conception occur. In individuals who do not conceive in a given cycle, the luteal gland regresses after approximately two weeks. Luteal regression may, or may not, play a role in the intraovarian

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regulation of subsequent folliculogenesis and selection of the next ovulatory or nonovulatory dominant follicle [35,205]. The new luteal gland can be recognized when the former “follicle walls” meet after follicle fluid is released and appears as two thickened slightly hypoechoic parallel tissue interfaces. The interface of the opposing former “walls” can be traced to the point of rupture at the outer edge of the ovary during the first week following ovulation. The CL becomes increasingly hypoechoic and thicker, reflecting increasing vascularity. Neovascularization of the CL begins immediately after evacuation of follicle fluid and is visualized with ultrasonography within 48–72 h as a vascular ring surrounding the developing luteal tissue. As the CL matures, the ring of vascularity becomes more prominent on color flow and power Doppler interrogation. The degree of vascular perfusion of the CL is also apparent when observing the gray scale image of the CL [104]. After the initial evacuation of the follicle fluid at ovulation, the CL may refill with a hypoechoic or specular fluid; many CL appear to have an echoic central cystic cavity [24,27,104,206]. Detection of fluid within the cavity of a CL has been interpreted as a normal physiologic event related to either leakage of blood from the vascular follicle wall into the CL lumen following follicle rupture or extravasation of blood into the central cavity during luteogenesis. Fluid may be observed immediately following ovulation and may subsequently decrease, remain, or increase in volume. The shape of a cystic area may therefore vary from a thin line or ovoid-shaped lumen to a round, cyst-like shape. The identity of a cystic CL can be confirmed as being distinct from a follicle by looking for the point of rupture on the external surface of the ovary during the first week after ovulation [104]. Once the point of rupture can no longer be identified, it may be difficult to differentiate between a cystic CL and an anovulatory follicle. Hence, the easiest time to identify a CL is shortly after ovulation. Color flow Doppler ultrasonography can confirm greater vascular flow within the CL wall compared to the vascularity expected for a preovulatory follicle. When blood fills the CL lumen, the cystic CL is regarded as a corpus hemorrhagicum (CH) [207]. Blood cells, clot, protein, and cellular debris in the fluid of the CH may lend a variable echotexture to the fluid-filled lumen [104,208]. CL can be observed in the ovaries throughout the luteal phase of the ovarian/menstrual cycle. The CL will regress with the onset of the next menses without conception or will persist through the first trimester of pregnancy. Following luteal regression, a corpus albicans may be visualized until the time of subsequent ovulation [104]. Occasionally, several corpora albicanthae may be observed from previous menstrual cycles. The location of small follicles surrounding the regressing corpora albicanthea may the influence their visualization.

Intraovarian effects of CL or CA on recruitment of follicles in their vicinity are unknown.

INSIGHTS REGARDING ANIMAL MODELS FOR THE STUDY OF FOLLICULOGENESIS IN WOMEN Research in humans, especially in elucidating our reproductive processes, is extremely limited by ethical and practical considerations. Therefore, we are required to make inferences regarding human reproduction from detailed studies in nonhuman primates, rodents, and domestic farm animal models. Nowhere has this limitation been more evident that in our work to elucidate ovarian function—especially that related to the dynamics of folliculogenesis—over time [37,64,75,95,126,209–212]. There are many similarities in folliculogenesis among various other mammals and humans; however, it is also critically important to recognize differences in physiology and the various experimental approaches that may be used in different species. The presence of endometrial exuviation at menses and luteal and follicular phases of equivalent length in women and nonhuman primates differ from the reproductive functions during the estrous cycles of domestic farm animals and laboratory animals. However, broad similarities in antral follicular dynamics exist irrespective of the differences. One of the most important similarities has been established by the use of frequent serial high-resolution ultrasonography and simultaneous endocrine profiling. The insights gained in detailed studies of folliculogenesis in mares and cattle have dramatically increased our broader understanding of follicular wave dynamics throughout the ovarian cycle. Ovarian follicular waves have now been described in every mammalian species in which an imaging-based approach has been used. The number of follicular waves that develop during each reproductive appears to be species-specific and is correlated with the length of the menstrual/estrous cycle. It is noteworthy that the growth phase of the dominant preovulatory follicle (defined as the time period from emergence of the recruited “cohort” or “wave” to ovulation) is comparable among humans, cows, and mares despite a proportionately longer luteal phase in cows and mares [41,46,64]. Detailed studies of dynamic ovarian function in humans will require frequent, perhaps as often as several times per day, blood sampling and ultrasonography depending on the specific hypothesis or function under examination. Studies of this nature in humans are difficult, and often unethical, to conduct. However, differences in folliculogenesis between humans and other mammals appear to be more in detail rather than in essence, and they may reflect differences in intrinsic physiology and/or differences in our ability to detect changes

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SUMMARY

in ovarian function with a single experimental approach [43,71,73,84,96,114,125,213,214]. The relatively large diameters of antral follicles in cows and mares, compared to monkeys, sheep, and rodents, provide greater feasibility for characterizing antral follicular dynamics ultrasonographically. Thus, at present, cows and mares may be the most appropriate models for the ultrasonographic study of antral follicular dynamics in women [64,95]. Two especially pertinent contributions of the equine model are expected to be in the elucidation of the origin and mechanisms underlying the development of major and minor waves and the mechanisms of elaboration of two vs three waves of folliculogenesis during the IOI [64,72,75,89]. One way to mitigate the invasiveness of experimental work in humans may be to focus on very tightly defined times in the estrous cycle of animal models and menstrual cycles of women based upon the direct juxtaposition of the endocrinologic and morphologic status of their follicle wave dynamics. This approach may allow inference of detailed mechanistic inquiry with a minimally invasive approach in human subjects [35,64,75,87,95,215,216].

FUTURE DIRECTIONS AND CLINICAL IMPLICATIONS We are awaiting for the arrival of noninvasive imaging technology that will allow a “virtual histology” approach to visualizing ovarian function in real time. Our understanding of the critical steps in folliculogenesis that occur below the limits of resolution of the current generation of ultrasound instruments is crucial to the development of safer, more effective contraception and to optimizing ovarian stimulation regimens. The transition from secondary follicles to early antral follicles would be a critical juncture for therapeutic intervention. Accurate depictions of the growth patterns of 2–3 mm follicles are also highly desirable. These small follicles are near the detection limit of our current ultrasound instruments and there is a great deal of error in detecting them. Much physiology that we do not yet understand appears to occur in very small follicles. For example, 1–3 mm follicles develop in a wavelike manner following surges in plasma concentrations of FSH waves in cattle [91]. Similar studies in women have not been conducted. It will be beneficial to enhance our understanding of the endocrine and paracrine mechanisms underlying recruitment of primary and secondary follicles into antral follicular waves, to elucidate the origins and actions of a third subtle, but significant, FSH peak and fluctuations in luteal phase estradiol in women with three follicle waves and to parse the mechanism of selection of dominant, preovulatory follicles. The antral follicle count (AFC) is used routinely in clinical assessments for various aspects of infertility

investigations and therapies. However, there is considerable variation in the ability to achieve consistent results. Similarly, small antral follicles may gain or lose fluid volume sufficient to alter their ability to be visualized due to changes in their physiologic status between assessments. Understanding the daily changes in AFC during the menstrual cycle has important implications for determining the most appropriate time to assess the number of detectable follicles as a predictor of response to ovarian stimulation or to interpret relationship of the AFC to the stage of the ovarian cycle [217–222]. The idea that the human ovarian cycles are based upon wave patterns of folliculogenesis is a cornerstone of the notion that ovarian stimulation therapies can be initiated at different times during the cycle. Initiation of ovarian stimulation during the luteal phase in women is supported by the observation that multiple dominant follicles were induced to grow following FSH administration in the early luteal phase of monkeys [223]. Similarly, good results have been obtained in studies where ovarian stimulation was initiated following lute-ectomy in cattle [224]. Identification and synchronization of the start of follicle wave emergence and initiation of ovarian stimulation have been used to optimize follicular responses in cattle and women with a history of poor response to treatment [225,226]. Further support for concept of multiple follicular waves is provided by clinical reports of successful luteal phase oocyte retrieval and oocyte retrieval followed by in vitro maturation as an optional procedure for urgent fertility preservation [227–229]. The existence of healthy antral follicles available for retrieval in the luteal phase was consistent with, and attributed to, the emergence of a new wave of follicle development in the luteal phase.

SUMMARY Research conducted over the past 60 years using histologic, endocrinologic, and ultrasonographic techniques has profoundly increased our understanding of ovarian folliculogenesis in women. Comprehensive review of the literature shows similar trends among studies and among species [35]. It now seems clear that multiple cohorts—also referred to as “waves”—of antral follicles are recruited (emerge) during the human menstrual cycle. Observations of ovarian follicular waves in women are comparable to those documented in several animal species; however, species-specific differences exist. Elucidation of endocrine and paracrine regulation of the interplay among follicles as they develop in the ovaries is important for understanding follicle-oocyte interactions and identifying noninvasive markers of the physiologic status of follicles predictive of oocyte competence and assisted reproduction outcomes.

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C H A P T E R

4 Development of the Mammalian Ovary and Follicles Katja Hummitzsch, Helen F. Irving-Rodgers, Jeff Schwartz, Raymond J. Rodgers Abbreviations Bmp2 Ctnnb1 DAX1 DAZL dpc E Emx2 Fog2 FOXL2 FST/ fst Gata4 GREL LGR5 Lhx9 Lypd6 Magi2 OCT3/4 PD PGC Rspo1 Sf1 SYPC3 VASA WNT4/ Wnt4 Wt1

mammals include the ovulation of oocytes and production of hormones that act to coordinate reproductive activity among organs. The endocrinology of the adult ovary is relatively unique as the regulation of hormone secretion by the ovary is determined by formation and regression of the follicles and corpora lutea. The diversity of cell types in the ovary serves a breadth of endocrine, oogenic, and other functions. The multiplicity of cell types dictates the necessity of precise cell-cell communication within the ovary for proper function. Much recent research in the ovary has focused on how cells are regulated and how they communicate with each other. It is therefore of increasing importance to understand the cell lineages and cell-fate decisions that occur cyclically and linearly during the functional life of the ovary. Since much of the research effort in this area has been directed toward humans and agricultural and laboratory species, we will mainly reference this literature. Much of the published research on fetal development of ovaries in primates and agricultural species relies extensively on timed histological observations of the developing ovary. Using this approach, recent research has been directed toward the identification and study of the epithelial and the stromal compartments and has documented the important role of mass migration of stromal cells during formation of the ovary. Studies of mice, in particular, have relied heavily on lineage tracing techniques and on manipulation of gene expression during development. Additionally, many studies of rodents have focused on comparisons between development of the female and male gonads. Although this is important, less is known about the systematic development of the ovary. Thus, the understanding of development of the ovary across species is somewhat disjointed. The origin of granulosa cells was always believed to be an epithelial cell, but over the years, three separate

bone morphogenetic protein 2 beta-catenin dosage-sensitive sex reversal, adrenal hypoplasia critical region, on chromosome X, gene 1 deleted in azoospermia-like day post coitum embryonic day (mouse development) empty spiracles homeobox 2 friend of GATA forkhead box L2 follistatin GATA binding protein 4 gonadal-ridge epithelial-like leucine-rich repeat-containing G-protein coupled receptor 5; also known as GPR49 or GPR67 Lim homeobox protein 9 LY6/PLAUR domain containing 6 membrane-associated guanylate kinase, WW and PDZ domain containing 2 octamer-binding transcription factor 4; also known as POU5F1 postnatal day primordial germ cell R-spondin 1 steroidogenic factor 1 synaptonemal complex protein 3 mouse vasa homologue, MVH or DEAD (Asp-Glu-Ala-Asp) box polypeptide 4, DDX4 Wingless-Type MMTV Integration Site Family, Member 4 Wilm’s tumor suppressor gene

Introduction The reproductive strategies that have evolved in different species have been the result of strong natural selection pressures. Thus, it is no surprise that a wide variation exists between species in the development and structure and function of female gonads. The key roles of ovaries in

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4. DEVELOPMENT OF THE MAMMALIAN OVARY AND FOLLICLES

Granulosa cells of rete ovarii origin (From regressing nephrons of the mesonephros)

(A)

Granulosa cells of surface epithelial origin

(B) Surface epithelium

Mesonephric tubules

Basal lamina

Fibre

Granulosa cell

Oogonia, oocyte

Stromal cell

Capillary

FIG. 1 Illustrations of earlier theories on key aspects of ovary formation. (A) The somatic epithelial granulosa cells were considered to arise from other epithelial cells of rete ovarii of the medulla [1], which in turn arose from the nephrons of the mesonephros. (B) The surface epithelial cells contribute somatic cells to the ovigerous cords, which in turn give rise to the somatic epithelial granulosa cells [2,3].

pathways were hypothesized. Initially, they were proposed to be derived from the nephrons of the mesonephros (Fig. 1; reviewed recently in Refs. [4, 5]). The mesonephros acts as a temporary kidney early in gestation and contributes cells to the developing gonads, differently between the male and female ([6–12]; for review see Refs. [13, 14]). In ovaries, mesonephric tubules are found in the hilum and medulla where they are referred to as the rete ovarii. They persist into adulthood. The hypothesis that rete ovarii give rise to granulosa cells was based on the close association of rete ovarii with oocytes [1,15]. This was further strengthened by the demonstration that the presence of rete ovarii correlated with onset of meiosis [16] and follicle formation [17]. Subsequently, it was suggested that cells derived from the ovarian surface epithelium give rise to the granulosa cells [2,3]. However, a more recent examination of bovine ovarian development suggested that granulosa cells are not derived from differentiated ovarian surface epithelial cells; rather, both the apical ovarian surface epithelium and the granulosa cells arise from a common precursor population of gonadal ridge epithelial-like (GREL) cells [18]. GREL cells, in turn, are derived from cells of the surface of the mesonephros [18], which replicated to form the gonadal ridge/ovarian primordium. This is the model that we present in this chapter. The origins and development of germ cells and the cumulus cells are covered in other chapters. This chapter will focus on the origins of somatic cells and the formation of structures of the ovary.

GONADAL (OR GENITAL) RIDGE FORMATION The sexually undifferentiated gonadal ridge forms as a thickening of the coelomic or surface epithelium in an anterior/posterior direction on the ventral side of the mesonephros. The mesonephros acts as transient fetal kidney in mammals and in females regresses around 55 days post coitum (dpc) in cow (term 283 d) [19] and 75 dpc in sheep (term 147 d) [3]. Gonadal ridge formation occurs 33 dpc in humans (term 266 d) [20], between 27 and 31 dpc in the cow [21], 23 dpc in sheep [22], 25 dpc in goat (term 150 d) [23], and embryonic day (E)10.3–10.4 in mice (term 20 d) [24] (Table 1). The initial thickening and full development of the gonadal ridge requires the expression of the transcription factor Gata4 (GATA-binding protein 4; [24]) in cooperation with Fog2 (Friend of GATA; [51]), Sf1 (steroidogenic factor 1), Wt1 (Wilm’s tumor suppressor gene), Lhx9 (Lim homeobox protein 9; reviewed in Ref. [52]), and Emx2 (empty spiracles homeobox 2; [53]). Conditional knockout of Gata4 in mice decreased proliferation of coelomic or surface epithelial cells before the thickening process of the gonadal ridges would normally occur and to reduced Sf1 and Lhx9 expression [24]. Homozygous null mutations in any of the other genes resulted in regression of the developing gonadal ridge in mouse embryos. The coelomic or surface epithelium of the mesonephros proliferates and undergoes phenotypical changes (Fig. 2A–C), giving rise to GREL cells in bovine [18],

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GONADAL (OR GENITAL) RIDGE FORMATION

TABLE 1

Comparison of Key Events During Ovarian Development Between Rodents, Primates, and Ruminants Rodents

Primatesa

Ruminants

Mouse: 18–22 days Rat: 21–23 days

Human: 40 weeks

Cow: 279–287 days Sheep: 144–152 days Goat: 148–152 days

Formation

Mouse: E10.3–10.4 [24] Rat: E12 [25]

Human: week 5 [20]

Cow: 27–38 dpc [19,21] Sheep: 23 dpc [22] Goat: 25 dpc [23]

Thickening of coelomic epithelium

Mouse [24] Rat [25]

Human [26,27]

Cow [19]

Basement membrane dissolution (below coelomic epithelium/mesonephric surface epithelium)

Mouse [24]

Human [26–28]

Cow [18,19]

Sexual differentiation

Mouse: E10.5–11.5 dpc [29] Rat: E14.5 [30]

Human: week 6–7 [20] Rhesus monkey: week 7–8 [31]

Cow: 40 dpc [21] Sheep: 28 dpc [22] Goat: 34 dpc [23]

PGC migration

Mouse: E9.5–11.5 [32]

Human: week 5 [20,33]

Cow: 23–31 dpc [21]

PGC arrival at genital ridge

Mouse: E10.5 [34]

Human: week 6 [33]

Cow: 32 dpc [21] Sheep: 23 dpc [3]

Germ cell differentiation

Synchronized in ovigerous cords; from anterior to posterior of ovary [34]

Zonal; medullar (mature) to proximal (immature) [18,31]

Zonal; medullar (mature) to proximal (immature) [35]

Mitosis

Rat: E14.5–18.5 [36]

Human: 2–7 months [36] Rhesus monkey: 7–19 weeks [31]

Cow: 50–150 dpc [37]

Germ cell nests/cysts

Mouse [34]

Human [38]

Cow [19]

Entry into meiosis

Mouse: E13.5 [39] Rat: E17–18 [40]

Human: week 11 [35] Rhesus monkey: 8 weeks [31]

Cow: 70–80 dpc [37] Sheep: day 55 [2] Goat: 55 dpc [23]

Waves of germ cell apoptosis

Mouse: E11.5–13.5 and E17.5–D9 [39] Rat: E17–20 [40]

Human: around week 22 [41]

Cow: 130–170 dpc [37] Sheep: 75–100 dpc [36]

Meiosis arrest

Mouse: E17.5-PD5 [34]

Gestational length

Genital ridge development

Oogenesis

Cow: 120–150 dpc [37]

Ovigerous cords Formation

Mouse: E13.5 [42] Rat: E13 [25]

Human: week 9–12 [28]

Cow: around 60 dpc [43] Sheep: 38–75 dpc [2] Goat: 36 dpc [23]

Connection with surface/no. separation from ovarian surface by basement membrane (early stage)

Mouse [42]

Human [28]

Cow [18,19] Sheep [4]

Breakdown

Mouse: E17.5 [39]

Human: week 20 [39]

Cow: 90–130 dpc [18,44]

Mouse: after birth [PD1–7, [45]] Rat: week 1 after birth [46]

Human: week 18–20 [28,47] Rhesus monkey: 20–23 [31]

Cow: around 90 dpc [44] Sheep: around 100 dpc [2] Goat: 90 dpc [23]

Follicle formation and activation Primordial follicle

Continued

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4. DEVELOPMENT OF THE MAMMALIAN OVARY AND FOLLICLES

Comparison of Key Events During Ovarian Development Between Rodents, Primates, and Ruminants—cont’d Rodents

Primates

Ruminants

Region of primordial follicle Formation

Mouse: medulla (first wave), cortex (second wave) [48] Rat: inner zone [46]

Inner cortex, close to medulla [28]

Inner cortex, close to medulla [2,18]

Primary follicle

Rat: PD5 [49]

Human: at 40 weeks all stages of folliculogenesis observed [50]b

Cow: 140 dpc [44] Sheep: 90 dpc [3]

Preantral follicle

Cow: 210 dpc [44] Sheep: 120 dpc [3]

Antral follicle

Rat: PD15 [49]

Cow: 240–285 dpc [19] Sheep: 135 dpc [3]

a

Literature about nonhuman primates with specific time points for developmental stages is rare. It mostly focuses on physiological aspects during pregnancy or on generational effects in disease animal models such as PCOS. Human fetal ovary samples from the third trimester are rare, and therefore the exact timing of the different follicle types after follicle activation is not available.

b

(A)

(B)

(C)

(D)

(F)

(E)

(G)

Surface epithelium

(H)

Germ cells:

GREL cell

Fibre

Granulosa cell

Capillary

Stromal cell

Basal lamina

Primordial Mitotic

Degenerative Meiotic

Oogonia Primary oocyte (arrested at prophase l)

FIG. 2 Schematic diagram of ovarian development. The development of the ovary commences at the mesonephric surface (A) in the location of the future gonadal ridge. Some mesonephric surface epithelial cells change phenotype into GREL cells (B). (C) The GREL cells proliferate and the basal lamina underlying the mesonephric surface epithelium breaks down. GREL cells continue to proliferate and PGCs (gray) migrate into the ridge between the GREL cells (D). Stroma from the mesonephros, including blood vessels, continues to expand and penetrates the ovary toward the ovarian surface. As it does so it branches and thus corrals proliferating oogonia and GREL cells into forming the ovigerous cords (E). The cords are surrounded by a basal lamina at their interface with stroma, but are open to the ovarian surface, and compartmentalization into cortex and medulla becomes obvious. Once the stroma has expanded to just below the surface the outermost GREL cells become partitioned on the surface separated from the underlying stroma by a basal lamina and then the GREL cells differentiate into the ovarian surface epithelial cells (F). Ovigerous cords are partitioned into smaller cords, commencing first at the interface with the medulla. Eventually, the cords are partitioned into follicles containing GREL cells that differentiate into granulosa cells and oogonia that become oocytes. The first primordial follicles appear in the inner cortex-medulla region, surrounded by a basal lamina (G). (H) Finally, the ovarian surface is covered by a simple epithelium. A tunica albuginea, densely packed with fibers, develops from the stroma below the surface epithelial basal lamina. Some primordial follicles become activated and commence development into primary and preantral follicles. Modified from Hummitzsch et al. [18] and incorporating information from Smith et al. [54].

I. THE OVARIAN FOLLICULAR APPARATUS: OPERATIONAL CHARACTERISTICS

OVIGEROUS CORD FORMATION

which start to express the cytoskeletal protein vimentin, as occurs with the subepithelial mesenchyme in the rat [25]. GREL cells express cytokeratins 18 and 19, plakophilin-2, and desmoglein-2 as do mesonephric surface epithelial cells [18]. Expression of cytokeratins 8, 18, and 19 has been found in the undifferentiated gonad in the mouse [42], whereas no cytokeratin 18 expression was detected in the rat [55]. Also during development, the basal lamina underlying the mesonephric surface epithelium where the gonadal ridge forms undergoes remodeling in a process controlled by Gata4 [24] as shown in mice. A similar incomplete basal lamina has been also observed in bovine ovarian development [19]. Presumably, this allows the primordial germ cells (PGCs) and mesenchymal or stroma cells from the mesonephros to penetrate between the proliferating surface epithelial cells of the developing gonadal ridge [18,26]. Mesonephric cell migration has been observed in the ovaries of human around 46 dpc [26] and sheep as early as 30 dpc (reviewed in Ref. [54]). This penetrating mesonephric stroma contains a vascular capillary bed, providing blood supply to the developing ovary. Thus, the early ovarian primordium contains two population of somatic cells; cells similar in appearance to the coelomic or surface epithelium, now termed GREL cells, and cells similar to blastemal cells of the mesonephros [26]. Based upon the description of their penetrating behavior, these blastemal cells of the mesonephros are the stromal cells from the mesonephros (Fig. 2D). PGCs, the carrier of genetic information of the next generation and the precursors for oogonia and oocytes, arise from the yolk sac (for review see [33,56–59]) and migrate through the primitive gut into dorsal mesentery and then laterally to the gonadal ridges. This process is regulated by the expression of stem cell factor and stromal-derived factor-1 by the gonadal ridge and surrounding mesenchyme (reviewed in Ref. [33]). In mice, PGCs start to migrate at E7.5 and enter the gonadal ridge at E10.5 ( [60], Table 1). In humans, the migration occurs in gestational week 5 and the PGCs arrive at week 6 (reviewed in Ref. [33]). PGCs begin to proliferate during the migration process. The colonization of the gonadal ridges by PGCs is followed by sex determination (reviewed in Ref. [61]) and subsequent differentiation into oogonia in the developing ovary (reviewed in Ref. [33]). The sex determination occurs in the human at week 6–7 [20], around 40 dpc in the cow [21], at 28 dpc in the sheep [22], after 25 dpc in the goat [62], at 23 dpc in the pig (term 114 d) [63], and around E10.5–11.5 in mice [29] (Table 1). The differentiation into an ovary is characterized by increased expression of Wnt4 (wingless-type MMTV integration site family, Member 4), Ctnnb1 (beta-catenin), Rspo1 (R-spondin 1), Foxl2 [Forkhead box 2, [64]], FST (follistatin), DAX1 (dosage-sensitive sex reversal, adrenal hypoplasia

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critical region, on chromosome X, gene 1), and meiosismarker SYCP3 (synaptonemal complex protein 3) [65]. Loss of expression of some of these genes, such as Foxl2 and Wnt4, results in female-to-male sex reversal (reviewed in Ref. [52]). PGCs migrate by amoeboid movement to the gonadal ridge during which they begin to proliferate. Once in the gonadal ridge, they cease migration and are classified as oogonia. Oogonia have a higher mitotic rate than PGCs and form germ cell nests. These oogonial nests undergo mitosis synchronously, and the oogonia are connected by intercellular bridges caused by cytokinesis that is incomplete at this stage [19,34,38]. Recently, it has been shown that the proliferation of germ cells is dependent upon the expression of Wt1 in somatic cells of the gonadal ridge [60]. Functional point mutation of Wt1 in mice resulted not only in a decrease of cell number from mitotic arrest and morphological change from an epithelial to mesenchymal phenotype in somatic cells but also in a decrease in germ cell numbers early in oogenesis (E10.5–12.5).

OVIGEROUS CORD FORMATION After formation of the gonadal ridge and sex differentiation, the stroma from the mesonephros penetrates further into the gonadal ridge/ovarian primordium, which at this stage is composed of GREL cells, PGCs, and oogonial nests [18]. This process creates areas of stroma, characterized by expression of COUP-TfiI [(also known as nuclear receptor subfamily 2 Group F Member 2 (NR2F2)] in stromal cells, alternating with areas of GREL cells/germ cells. This in effect produces the ovigerous cords composed of GREL cells and germ cells, and the cords are therefore initially “open” to the surface of the ovary as no distinct surface epithelium underlain by a basal lamina has been established at this stage [2,18,19,28] (Fig. 2E). The ovigerous cord formation occurs gradually from inside of the primordium toward the outer zone in line with the branching of the stroma as it penetrates toward the surface of the ovary. The penetrating stroma has been observed previously in human [50] and described as “cell streams” in sheep [3]. Importantly, the ovigerous cords are separated from the stromal compartment by a continuous basal lamina [3,18,19,28], composed of subunits of laminin 111 and collagens type IV and type XVIII, perlecan, and nidogens 1 and 2. The somatic GREL cells inside the ovigerous cords strongly express FOXL2, a marker of granulosa cells (the role of FOXL2 in the ovary is discussed in Ref. [66]). Furthermore, the GREL cells still express cytokeratins 18 and 19 [18,67], plakophilin-2, and desmoglein-2 [18]. Expression of cytokeratins 8 and 16–19 has also been identified in the somatic cells of the

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ovigerous cords of the fetal rat ovary [25]. The entry of oogonia into meiosis is first initiated in oogonia at the base of the ovigerous cords closest to the medulla and is triggered by retinoic acid, which is synthesized by aldehyde dehydrogenases in the mesonephros (Aldh1a2/3) and in the ovary (Aldh1a1) [68].

REGIONALIZATION—CORTEX AND MEDULLA The formation of ovigerous cords results in regionalization of the developing ovary into cortex and medulla in human [69], bovine [18], sheep [2], and goat [23]. In mice, a morphological regionalization can be observed close to birth at day E18.5 [70], but at a molecular level, the cortex and medulla can be distinguished as early as E12.5 by expression of Bmp2 (bone morphogenetic protein 2, [70]), Lypd6 (LY6/PLAUR Domain Containing 6), and Magi2 (membrane-associated guanylate kinase, WW and PDZ domain containing 2, [71]) in the cortex and Wnt4 and Fst [70] in the medulla. In general, the medulla is relatively much smaller in mice than humans and other species studied, and in the adult mouse ovary, it is even less obvious (reviewed in Ref. [72]). The cortex contains the ovigerous cords early in development and a vascularized stroma. Proliferation of oogonia continues to occur in the outer cortex, and maturation into oocytes and formation of first follicles happen at the interface of inner cortex and medulla (Fig. 2G). The medulla is characterized by lymphatic and large blood vessels, connective tissue, and remnants of mesonephric tubules (rete ovarii) [19,50]. The greatest number of Ki67-positive stromal cells occurs mainly in the medulla between 120 and 180 dpc in bovine [19].

polypeptide 4 (DDX4) [18,35]). Since there is a basal lamina at the interface of the penetrating stroma and the ovigerous cords, the GREL cells are the only somatic cells inside the cords, rendering them to be the precursor of the granulosa cells of follicles [18]. Less clear is whether GREL cells are identical to pregranulosa cells of primordial follicles or if GREL cells differentiate into pregranulosa cells upon follicle formation. The factors that initiate any transition are also unknown. As development progresses, significant proportions of oogonia undergo apoptosis, with the oogonial nests being reduced to individual oocytes arrested at the pachytene stage of meiosis I. These are surrounded by a finite number of GREL cells to form the primordial follicles (Fig. 2F). The basal lamina that thus far separated the ovigerous cords from the surrounding stroma now surrounds individual follicles. The number and potential quality of follicles with which the ovary is endowed, are ultimately determined by the complex interactions between the germ cell, GREL cells, and the stroma. Follicle formation occurs in human, ruminants (cow, sheep, goat), and pig before birth, whereas in the less precocious mice and other rodents, follicle formation starts shortly after birth (Table 1). Reports of the first primordial follicles in the human vary from as early as week 11.5 [80] to as late as between 16 and 20 weeks of gestation [47,81]. In cattle primordial follicles appear at 90 dpc [44], around 100 dpc in sheep [2], between 60 and 90 dpc in goat [82], at 70–90 dpc in pig [83], and in mice between postnatal days (PD)1–7 [45]. The large disparity in the emergence of primordial follicles observed, even in the mouse, suggests that this is a real biological variation, and not just due to experimental methodological differences.

OVIGEROUS CORD BREAKDOWN AND FOLLICLE FORMATION

ESTABLISHMENT OF PRIMORDIAL FOLLICLE RESERVE

Later in gestation, the stroma continues to penetrate the cortical region and the ovigerous cords, partitioning them into smaller groups of germ cells and GREL cells (Fig. 2F). The gradual breakdown of ovigerous cords proceeds from the interface between the medulla and the cortex and extends to the periphery of the cortex as the stroma expands toward the surface, as has been detected in fetal human [35,50,73], cattle [18,74,75], sheep [2,36,76], mouse [77], and postnatal rat ovaries [78,79]. It reflects the same gradual pattern reported in the maturation process of oogonia/oocytes as observed by expression of germ cell markers temporally from OCT3/4 (octamer-binding transcription factor 4; also known as POU5F1), DAZL (deleted in azoospermialike), and then to VASA (also known as mouse vasa homologue, MVH or DEAD (Asp-Glu-Ala-Asp) box

At the early stages of ovarian development, oogonia are highly proliferative that results in a peak in germ cell numbers with 6,800,000 between gestational weeks 16–20 in human [84], 2,700,000 at 110 dpc in cow [37], 850,000 at 75 dpc in sheep [36], approximately 15,000 at E15–20 in mice [41], in the fifth gestational month in rhesus monkey [31], and at E18.5 in rat [84]. These numbers then reduce drastically because of declining germ cell proliferation rates and increasing cell death. The total number of germ cells decreases by 80%–90% between 5 months and birth in human [84], 75–100 dpc in sheep [36], and 130–170 dpc in cattle [37], and by 60% between E18.5 and PD2 in rat [84]. It has been shown for rat fetal and postnatal ovaries and human fetal ovaries that there are three waves of germ cell degeneration [84]. The first wave includes oogonia undergoing mitosis (shortly before meiotic entry),

I. THE OVARIAN FOLLICULAR APPARATUS: OPERATIONAL CHARACTERISTICS

FORMATION OF THE OVARIAN SURFACE EPITHELIUM

the second waves affects oocytes at pachytene stage, and the third concerns oocytes at diplotene stage [84]. Two waves of germ cell death have been reported in mice. The first occurs during the period of meiosis at E13.5–15.5 and the second wave during follicle formation at E17.5–PD 1 [85]. The main mechanism for germ cell death is apoptosis mediated by gene products of the BCL2 family members (reviewed in Ref. [54]). Recent studies in mice suggest also an involvement of autophagy in germ cell death as a response to nutritional stress around birth. In addition, some oocytes on the surface of the ovary are lost by germ cell extrusion [86]. Reasons for the elimination of germ cells are chromosomal abnormalities (failure of mitosis and meiosis), defective mitochondrial genomes, insufficient pregranulosa cells, and degeneration of oocytes during restructuring of ovigerous cords and cysts into follicles (for review see Ref. [34, 41]).

FOLLICLE ACTIVATION Activation of primordial follicles and subsequent differentiation into primary, preantral, and antral follicles can be observed late in gestation in the cow [19] and sheep [3] (Table 1). Follicles in the mature stages undergo atresia before birth [19], such that only primordial follicles remain in the ovary. Similar events occur in mice where two classes of primordial follicles have been identified—medullary primordial follicles which will be activated shortly after birth and go through folliculogenesis fast before dying by apoptosis/follicular atresia in the first 3 months after birth and cortical follicles which will be activated through adulthood (reviewed in Ref. [48]). The medullary follicles contain granulosa cells arising from precursor cells that expressed Foxl2 in the fetal ovary, whereas granulosa cells from cortical follicles are derived from Lgr5 (Leucine-rich repeat-containing G-protein coupled receptor 5; also known as GPR49 or GPR67)-positive cells, which were located in the cortex and at the ovarian surface. Mork et al. [87] concluded that the two distinct granulosa cell population might still descend from one progenitor line that would be compatible with the GREL cell model described in the cow [18].

FORMATION OF THE OVARIAN SURFACE EPITHELIUM Previous literature implied that the mature ovarian surface epithelium originates from the mesodermderived epithelial layer lining the intraembryonic coelom and the area where the gonad is formed. The gonadal blastema or ovarian primordium is partly formed by proliferation of this surface layer (reviewed in Ref. [88]).

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Interestingly, when the ovarian primordium first forms, it lacks a subepithelial basal lamina, except at the base of the ovary where it arises and protrudes from the mesonephros (Fig. 2D) (cow [18,19], human [28], and mouse [42]). Thus, the base or hilum of the ovary is mesonephros and its epithelium remains intact while the remainder of the primordium develops. All cells on the surface of fetal mice ovaries, except those at the base, express Lgr5 at E16.5 [89]. This may contribute to the observation that the hilum or base of postnatal mice ovaries is richer in stem cells with greater oncogenic potential than other locations on the ovarian surface [90]. The absence of a mature surface epithelium during early ovarian development is further supported by the observation that surface epithelial cell expression of estrogen receptor 1 (also known as ERα), a characteristic of the adult ovarian surface epithelial layer, does not occur until at least gestational day 55 in sheep [91] and day 75 in cattle [43]. The early ovarian primordium is composed of GREL cells that differentiate from the mesonephric surface epithelium, together with germ cells, and stroma penetrating from the mesonephros. GREL cells lining the outside of the developing ovary are tightly connected by adherens junctions to form a protective cover [18]. At the stage of ovigerous cords, the GREL cells nearest the surface appear to express cytokeratin 19 [19] and the desmosomal proteins plakophilin-2 and desmoglein-2 more strongly than the GREL cells closer to the medulla [18]. Similar zonal expression has been observed in mouse ovaries, with only cells in the outer cortical zone being positive for cytokeratin 19 [42]. Through the branching and expansion of stroma into the cortical area and lateral extension below the ovarian surface, a basal lamina forms under the GREL cell layers at the surface. Thus, the GREL cells on the surface begin to differentiate into surface epithelial cells. A well-defined basement membrane can be observed around the fifth month of human gestation (reviewed in Ref. [88]) and around 150 dpc in bovine [18]. At this point, the surface epithelium is still multilayered and becomes a single layer between 180 and 200 dpc in bovine [18,19] as a result of increased apoptosis from 165 dpc onward [19]. The current model of formation of the ovarian surface epithelium and granulosa cells of follicles has both cell types derived from precursor GREL cells. Support for the concept that both, granulosa and surface epithelial cell types, might arise from one precursor cell [18] comes from the common morphology described in bovine fetal ovaries of elongated columnar cells with elongated central nuclei and dark eosinophilic cytoplasm [19]. Another important consideration is that there are two developmental origins of ovarian surface epithelial cells. The initial surface epithelial cells covering the major apical portion of the developing ovary are derived from the GREL cells, which in turn were originally derived from the mesonephric surface epithelial cells. However, the

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hilum or base of the ovary is directly derived from mesonephros and its epithelium is a classic epithelium that remains as such during development of the ovary. The implications of two apparently distinct developmental pathways are not known but are possibly related to the higher onogenicity and stemness of epithelial cells at the hilum versus the remainder of the apically situated surface epithelial cells [92].

FORMATION OF THE OVARIAN TUNICA ALBUGINEA The tunica albuginea is a nonvascularized [93], thick fibrous-rich layer, composed mainly of structural collagens and other extracellular proteins (collagen type I, decorin, versican), located below the ovarian surface [18,94,95]. The ovarian tunica albuginea is not as thick as that of the testis and forms late during development, after the ovarian surface epithelium has been established. In cattle, this occurs between 240 and 285 dpc [19] and in rat around E17 [25]. The tunica albuginea appears to have some degree of zonation and is variable in thickness from one location to another in the ovary [95]. The tunica albuginea originates from the mesonephric stroma that penetrates the ovary in “cell streams” that extend to just below the surface as described previously (Fig. 2) [18]. Unlike the other stromal compartments within the ovary, the tunica albuginea and the outer cortex containing primordial follicles are avascular. The factors initiating the changes in the cortical stroma resulting in formation of the tunica albuginea are not known. Given the proximity to the surface epithelium, it is possible that factors from the surface epithelial cells influence the adjacent stromal cells to develop into the fibrous tunica. Aberrant changes to the tunica albuginea have been observed in human conditions such as polycystic ovary syndrome (PCOS), where ovaries exhibit a substantially thicker tunica albuginea containing more collagen and also have increased thicknesses of cortical and subcortical stroma [96]. This might negatively influence folliculogenesis and ovulation.

FORMATION OF THE THECA INTERNA AND EXTERNA The first thecal cells can be identified at the follicular preantral stage and therefore during late gestation in human [50], bovine [19], and sheep [3], but only after birth in mouse and rat [97]. Thecal layers form close to the basal lamina surrounding the membrana granulosa of the follicle and they start to differentiate into a theca interna and theca externa during follicle growth and antrum formation. The outer layer, the theca externa is

composed of fibroblasts, collagen fibers, larger venules, lymphatic vessels, nerve fibers, and cells with contractile filaments, whereas the theca interna contains specialized steroidogenic cells, fibroblasts, immune cells, and capillaries. These steroidogenic thecal cells synthesize androgen precursors required by granulosa cells for estradiol production. A role of the theca externa has not been identified, but it may play a role in follicular fluid expulsion during ovulation (reviewed in Ref. [98]). Steroidogenic thecal cells most likely arise from stem cells [99] within the stroma and indeed a potential stem cell niche in the theca has previously been identified [100]. Certain chondroitin/dermatan sulfate epitopes (antibodies 7D4, 3C5, and 4C3), which marked stem cell niches in other tissues, have been detected in the ovary. A study in pigs identified an alkaline phosphate-positive cell population in cultured thecal cells from small antral follicles [101]. In addition to the mesenchymal surface markers CD29, CD44, and CD90, the cells also expressed the pluripotency marker SOX2. Furthermore, these cells showed multipotency potential by differentiating into osteocytes, adipocytes, and oocyte-like cells. Recently in mouse, it was shown that Indian hedgehog (Ihh) and Desert hedgehog (Dhh) expression in granulosa cells is required for the formation of thecal cells, which express the downstream target Gli [102]. Among other effects, double-knockout of Ihh and Dhh results in disruption of folliculogenesis at the preantral stage and fewer CYP17A1-expressing cells in the ovarian mesenchyme. Furthermore, during gestation, Gli-positive cells are only found in the mesonephros, and it appears that Gli-positive cells from the mesonephros are the origin of stem cells for thecal lineage.

EARLY FORMATION OF THE VASCULATURE Vascularization of the developing ovary is less pronounced and occurs later than in the developing testis, making the degree of vascularization one of the earliest distinguishing morphological features between the female and male gonads [103]. When the stroma penetrates into the gonadal ridge/ovarian primordium, it contains endothelial cells assembled into mature capillaries surrounded by a subendothelial basal lamina as observed in human [28] and bovine fetal ovaries [18]. Blood vessels in human fetal ovaries express prokineticin receptor 1, which is involved in angiogenesis later in gestation (14–19 weeks; [104]). A complex vascular network, expressing PECAM1 and calveolin-1, has been observed early (E13.5) in the developing mouse ovary [105]. Disruption of calveolin-1 in mice resulted in decreased angiogenesis [106]. Furthermore, cells associated with vasculature are in the stromal compartment between

I. THE OVARIAN FOLLICULAR APPARATUS: OPERATIONAL CHARACTERISTICS

REFERENCES

germ cell nests between E14.5 and 18.5 and they express MafB (MAF BZIP Transcription Factor B) [107], which has been recently shown to be a regulator of endothelial cell sprouting [108]. Thus, the initial capillary network of the ovarian cortex is not likely formed by vasculogenesis but rather by the sprouting or splitting forms of angiogenesis [109], allowing expansion of the existing capillary network derived directly from vasculature in the mesonephros. This appears to also happen in mouse where is was observed that “few endothelial cells crossed the border between the mesonephros and the XX gonad” [110] and “the XX gonad recruits vasculature by a typical angiogenic process” [103]. Lymphatic vessels are visible in the hilum area/ medulla of the developing human ovary at 15 weeks [111], but in the mouse they only appear around PD10, the time when the first wave of growing follicles become estrogenic [112]. As the follicles continue to grow, highly branched lymphatic vessels are recruited to the theca and stromal layers around each follicle, resulting in the establishment of the ovarian lymphatic network [113,114]. Subsequently, the lymphatic network is remodeled to accommodate the growth of each new follicle wave throughout the reproductive life span [113,114]. Neolymphangiogenesis around developing follicles is prevented by blockade of VEGFR3 (vascular endothelial growth factor receptor 3) signaling, causing a reduction in follicle viability and hormone secretion [115].

CONCLUSIONS The latest model of ovarian development presented, while elegant in its simplicity, has been developed substantially only from timed histological observations of developing ovaries. Further study is indicated of how GREL cells develop into either surface epithelial cells or granulosa cells, and perhaps any influence of the oogonia/oocytes in this process. The developmental origins of surface epithelial cells at the base or hilum of the ovary differ from those on the rest of the ovary. Cells from these regions have been found to differ in stemness and oncogenicity. Given the recent new hypotheses on the cellular origins of ovarian cancer, this discovery warrants further investigations. The mesonephros-derived stroma penetrates into the developing ovary, branching as it does so. It is likely that the movement of the stroma is responsible for the physical formation of ovigerous cords, surface epithelium, and follicles. The penetrating stroma also brings with it the initial vascular supply of the ovary. Additional research is now needed to interrogate and refine these concepts further to enable a fuller understanding of the development of the ovary and its follicles.

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SRY transcript and developmental expression of genes involved in sexual differentiation. Int J Dev Biol 1996;40(3):567–75. Pailhoux E, Vigier B, Vaiman D, Servel N, Chaffaux S, Cribiu EP, et al. Ontogenesis of female-to-male sex-reversal in XX polled goats. Dev Dyn 2002;224(1):39–50. Hu YC, Okumura LM, Page DC. Gata4 is required for formation of the genital ridge in mice. PLoS Genet 2013;9(7)e1003629. Frojdman K, Paranko J, Virtanen I, Pelliniemi LJ. Intermediate filament proteins and epithelial differentiation in the embryonic ovary of the rat. Differentiation 1993;55(1):47–55. Wartenberg H. Development of the early human ovary and role of the mesonephros in the differentiation of the cortex. Anat Embryol (Berl) 1982;165(2):253–80. Gruenwald P. The development of the sex cords in the gonads of man and mammals. Am J Anat 1942;70(3):359–97. Heeren AM, van Iperen L, Klootwijk DB, de Melo Bernardo A, Roost MS, Gomes Fernandes MM, et al. Development of the follicular basement membrane during human gametogenesis and early folliculogenesis. BMC Dev Biol 2015;15:4. Hacker A, Capel B, Goodfellow P, Lovell-Badge R. Expression of Sry, the mouse sex determining gene. Development 1995;121(6): 1603–14. Mittwoch U, Delhanty JDA, Beck F. Growth of differentiating testes and ovaries. Nature 1969;224:1323. Baker TG. A quantitative and cytological study of oogenesis in the rhesus monkey. J Anat 1966;100(Pt 4):761–76. Anderson R, Copeland TK, Sch€ oler H, Heasman J, Wylie C. The onset of germ cell migration in the mouse embryo. Mech Dev 2000;91(1):61–8. De Felici M. Origin, migration, and proliferation of human primordial germ cells. In: Coticchio G, editor. Oogenesis XII. London: Springer-Verlag; 2013. p. 364. Pepling ME. From primordial germ cell to primordial follicle: mammalian female germ cell development. Genesis 2006;44 (12):622–32. Anderson RA, Fulton N, Cowan G, Coutts S, Saunders PT. Conserved and divergent patterns of expression of DAZL, VASA and OCT4 in the germ cells of the human fetal ovary and testis. BMC Dev Biol 2007;7:136. McNatty KP, Smith P, Hudson NL, Heath DA, Tisdall DJ, WS O, et al. Development of the sheep ovary during fetal and early neonatal life and the effect of fecundity genes. J Reprod Fertil Suppl 1995;49:123–35. Erickson BH. Development and radio-response of the prenatal bovine ovary. J Reprod Fert 1966;10:97–105. Motta PM, Makabe S, Nottola SA. The ultrastructure of human reproduction. I. The natural history of the female germ cell: origin, migration and differentiation inside the developing ovary. Hum Reprod Update 1997;3(3):281–95. Rosario R, Adams IR, Anderson RA. Is there a role for DAZL in human female fertility? Mol Hum Reprod 2016;22 (6):377–83. Wartenberg H, Ihmer A, Schwarz S, Miething A, Viebahn C. Mitotic arrest of female germ cells during prenatal oogenesis. A colcemid-like, non-apoptotic cell death. Anat Embryol (Berl) 2001;204(5):421–35. Kerr JB, Myers M, Anderson RA. The dynamics of the primordial follicle reserve. Reproduction 2013;146(6):R205–15. Appert A, Fridmacher V, Locquet O, Magre S. Patterns of keratins 8, 18 and 19 during gonadal differentiation in the mouse: sex- and time-dependent expression of keratin 19. Differentiation 1998; 63(5):273–84. Garverick HA, Juengel JL, Smith P, Heath DA, Burkhart MN, Perry GA, et al. Development of the ovary and ontongeny of mRNA and protein for P450 aromatase (arom) and estrogen

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[104] Eddie SL, Childs AJ, Kinnell HL, Brown P, Jabbour HN, Anderson RA. Prokineticin ligands and receptors are expressed in the human fetal ovary and regulate germ cell expression of COX2. J Clin Endocrinol Metab 2015;100(9):E1197–205. [105] Bullejos M, Bowles J, Koopman P. Extensive vascularization of developing mouse ovaries revealed by caveolin-1 expression. Dev Dyn 2002;225(1):95–9. [106] Morais C, Ebrahem Q, Anand-Apte B, Parat MO. Altered angiogenesis in caveolin-1 gene-deficient mice is restored by ablation of endothelial nitric oxide synthase. Am J Pathol 2012;180(4): 1702–14. [107] Maatouk DM, Mork L, Hinson A, Kobayashi A, McMahon AP, Capel B. Germ cells are not required to establish the female pathway in mouse fetal gonads. PLoS One 2012;7(10)e47238. [108] Jeong H-W, Hernández-Rodríguez B, Kim J, Kim K-P, EnriquezGasca R, Yoon J, et al. Transcriptional regulation of endothelial cell behavior during sprouting angiogenesis. Nat Commun 2017;8(1):726. [109] Kovacic JC, Moore J, Herbert A, Ma D, Boehm M, Graham RM. Endothelial progenitor cells, angioblasts, and angiogenesis—old terms reconsidered from a current perspective. Trends Cardiovasc Med 2008;18(2):45–51.

[110] Brennan J, Karl J, Capel B. Divergent vascular mechanisms downstream of Sry establish the arterial system in the XY gonad. Dev Biol 2002;244(2):418–28. [111] Kleppe M, Kraima AC, Kruitwagen RFPM, Van Gorp T, Smit NN, van Munsteren JC, et al. Understanding lymphatic drainage pathways of the ovaries to predict sites for sentinel nodes in ovarian cancer. Int J Gynecol Cancer 2015;25(8):1405–14. [112] Brown HM, Dunning KR, Robker RL, Pritchard M, Russell DL. Requirement for ADAMTS-1 in extracellular matrix remodeling during ovarian folliculogenesis and lymphangiogenesis. Dev Biol 2006;300(2):699–709. [113] Brown HM, Robker RL, Russell DL. Development and hormonal regulation of the ovarian lymphatic vasculature. Endocrinology 2010;151(11):5446–55. [114] Svingen T, Francois M, Wilhelm D, Koopman P. Threedimensional imaging of Prox1-EGFP transgenic mouse gonads reveals divergent modes of lymphangiogenesis in the testis and ovary. PLoS One 2012;7(12)e52620. [115] Rutkowski JM, Ihm JE, Lee ST, Kilarski WW, Greenwood VI, Pasquier MC, et al. VEGFR-3 neutralization inhibits ovarian lymphangiogenesis, follicle maturation, and murine pregnancy. Am J Pathol 2013;183(5):1596–607.

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C H A P T E R

5 Organization of Ovarian Steroidogenic Cells and Cholesterol Metabolism Jerome F. Strauss, III Department of Obstetrics and Gynecology, Virginia Commonwealth University, Richmond, VA, United States

INTRODUCTION

olefinic bonds; and the addition of hydroxyl functions, proceeding invariably from cholesterol through the pregnane (21 carbon backbone), androstane (19 carbons), and finally, estrane (18 carbons) families, although some have argued that shortcuts do exist. Specific cell types can accomplish several of these sequential steps, but rarely can they generate an estrogen from starting from cholesterol. Indeed, the requirement for cooperative efforts by two different tissues or cell types is a characteristic feature of estrogen biosynthesis. This joint effort enables the modulation of both androgen and estrogen production by factors that independently influence the cells involved in precursor synthesis, in addition to the cell type in which the final step of estrogen synthesis, aromatization, occurs.4 This collaboration is exemplified by estradiol synthesis in ovarian follicles, where luteinizing hormone (LH) acts on the theca cells to stimulate production of androgen precursors and follicle-stimulating hormone (FSH) acts on granulosa cells to stimulate aromatization of theca androgens into estrogens.

The ovary is the primary source of the two main female reproductive hormones, estradiol and progesterone. It also produces androgens and other bioactive sterols that function locally to regulate metabolism and possibly gamete function. These steroid hormones have endocrine, paracrine, and intracrine roles. In its endocrine role, estradiol triggers female sexual maturation and sustains the reproductive tract. Progesterone promotes differentiation and maintenance of cells and tissues that have been preprogrammed by estradiol. These hormones also act not only within the ovary in paracrine signaling but also within the cells that produce them (intracrine actions). In addition to sex steroid hormones, the ovary produces sterol-derived molecules implicated in oocyte maturation (meiosis-activating sterols) and the control of cellular cholesterol homeostasis (hydroxycholesterols). This chapter reviews the essential components of sterol metabolism and the steroidogenic machinery in the ovary, focusing on the initial steps in steroid hormone biosynthesis, the acquisition of the steroid hormone precursor, cholesterol, and its metabolism into pregnenolone, the first committed step in synthesis of steroid hormones.

ACQUISITION, STORAGE, AND TRAFFICKING OF CHOLESTEROL Steroid-producing cells have structural features that enhance their ability to obtain and store cholesterol for use in hormone synthesis (Fig. 1).2,6 Unlike protein hormone-producing cells, steroidogenic cells do not store prefabricated hormone; they synthesize hormone on demand from cholesterol that has been acquired from the plasma, synthesized de novo, or stored in membranes, or as sterol esters in lipid droplets. Because of cholesterol’s limited solubility in the cytosol, multiple proteins are required to move it efficiently from one cellular compartment to another either acting as transport

ORGANIZATION OF STEROIDOGENIC TISSUES AND CELLS The steroidogenic machinery is organized into compartments at the organ, cellular, and subcellular levels. This compartmentalization has important implications for the physiological control of hormone production.1–3 The synthesis of steroid hormones involves a series of sequential modifications of cholesterol that result in the removal of the side chain of cholesterol; alterations in

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SR-B1 LDL

Sterol efflux

LDL receptors Coated vesicle

Coated pit

HDL

ABCA1

Recycled LDL receptor

Cholesterol esters

P450scc SCP2 NPC1

NPC2

Free cholesterol

Free cholesterol STARD3

Acid Late lipase endosome

De novo cholesterol synthesis Acetyl-CoA

SCP2

STARD3 SOAT 1

Lysosome

STARD1

NCEH1/ LIPE

Cholesterol esters NCEH1/LIPE

HMG-CoA reductase

Perilipins Lipid droplet

Endoplasmic reticulum

FIG. 1 The acquisition, storage, and trafficking of cholesterol in steroidogenic cells. ABCA1, ATP-binding cassette transporter A1; FFA, free fatty acid; GRP78, glucose regulatory protein 78; HDL, high-density lipoprotein; HMG-CoA, 3-hydroxy-3-methylglutaryl-coenzyme A; LDL, low-density lipoprotein; LIPE, hormone-sensitive lipase; NCEH1, neutral pH cholesterol ester hydrolase; SR-B1, scavenger receptor type B; GRP78, glucoseregulated protein 78; STARD1, steroidogenic acute regulatory protein; STARD3; (steroidogenic acute regulatory protein)-related lipid transfer domain 3; STARD4; STARD5; SCP2, sterol carrier protein 2; SOAT1, sterol-O-acyltransferase-1; VDAC2, voltage-dependent anion channel 2. Used with permission from Elsevier.5

proteins or by creating contact points between membranes that facilitate movement of cholesterol from sterol-rich donor membranes to sterol-poor acceptors.

PLASMA MEMBRANE CHOLESTEROL The plasma membrane has the highest content of free cholesterol of all cellular membranes (60%–90% of cellular fee cholesterol). It is derived from plasma lipoproteins and de novo sterol synthesis. The plasma membrane sterol pool is not static, it exchanges with plasma lipoprotein free cholesterol, and regularly cycles through the cell and back to the plasma membrane. During this cycling process, cholesterol can be diverted for use in steroid hormone synthesis or esterified and deposited in lipid droplets for future use.

PLASMA LIPOPROTEINS: THE LDL AND HDL PATHWAYS The access of certain plasma lipoproteins to certain ovarian compartments is limited. For example, lowdensity lipoproteins (LDLs) and very low-density

lipoproteins (VLDLs) are large particles (>1 million molecular weight), and they cannot penetrate the blood-follicle barrier to reach granulosa cells prior to ovulation. Thus, granulosa cells of the preovulatory follicle must rely on other sources of cholesterol for hormone synthesis. It is only with neovascularization of the ovulated follicle that follicular cells can gain access to LDL and VLDL. Lipoprotein-gathering receptors of the LDL receptor family (e.g., LDL receptors, LDL receptorrelated protein, VLDL receptors) are located on the numerous microvilli that project from the plasma membrane of cells that produce large quantities of steroid hormone (e.g., luteal cells).2 These receptors mediate lipoprotein internalization by an endocytic mechanism that ultimately delivers the lipoproteins to lysosomes where the apolipoproteins are degraded and the lipoprotein-carried cholesterol esters are hydrolyzed to release free cholesterol. Stimulation of ovarian steroidogenic cells with gonadotrophic hormones increases the number of LDL receptors on the cell surface and also accelerates the rate of LDL endocytosis and degradation. Several key proteins are involved in the availability of cholesterol from lipoproteins internalized by the LDL receptor family. These include lysosomal acid lipase, encoded by the LIPA gene. Severe acid lipase deficiency,

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RELATIVE ROLES OF PLASMA LIPOPROTEINS AND DE NOVO SYNTHESIS IN SUPPLYING CHOLESTEROL FOR STEROIDOGENESIS

Wolman’s disease, is associated with lysosomal accumulation of cholesterol esters and triglycerides, which can lead to damage of steroidogenic cells and compromised hormone production.2 Free cholesterol is released from lysosomes by a system of sterol-binding proteins encoded by genes (NPC1, NPC2) that when mutated cause the cholesterol storage disorder, Neiman-Pick Type C disease. NPC2, a soluble protein in the lysosome, delivers free cholesterol to NPC1, a membrane-associated cholesterol-binding protein that controls sterol efflux from the lysosomes. Other sterol-binding proteins, including sterol carrier protein-2 (SCP2), steroidogenic acute regulatory protein (STAR)-related lipid transfer (START) domain protein 3 (STARD3), also known as metastatic lymph node 64 protein (MLN64), and STARD4 may participate in the intracellular trafficking of cholesterol to various organelles, including from endosomes to the endoplasmic reticulum, and ultimately to the mitochondria.2,7–9 In steroidogenic glands regulated by gonadotropic hormones, the delivery of cholesterol substrate to the ultimate destination, the mitochondrial cholesterol side-chain cleavage enzyme, is mainly effected by the steroidogenic acute regulatory protein (StAR or STARD1).2 High-density lipoproteins (HDLs) can also provide cholesterol for hormone synthesis by a pathway that differs from the “LDL pathway.”10 Unlike LDL, HDL being a smaller particle can penetrate into the follicle and accumulate in follicular fluid. Receptors for HDL (scavenger receptor type B, class 1 [abbreviated SR-B1]) are located in closely apposed microvilli that form “microvillar channels” in which HDL particles are lodged.3 Endothelial lipases, including hepatic lipase or endothelial cellderived lipases, may facilitate uptake of the HDL-carried sterols by steroidogenic cells, including the selective uptake the HDL sterol esters. Like LDL receptor expression, SR-B1 expression is upregulated in response to trophic stimulation, facilitating the usage of HDLdelivered substrate. HDL-carried free cholesterol and cholesterol esters are selectively internalized by SR-B1, leaving the HDL apolipoproteins on the cell surface. The internalized HDL cholesterol esters are then cleaved, by a cytosolic, neutral pH optimum sterol esterase (also referred to as hormonestimulated lipase encoded by LIPE) or neutral cholesterol ester hydrolase, encoded by NCEH1), releasing free cholesterol.6,10

DE NOVO CHOLESTEROL SYNTHESIS De novo synthesis of cholesterol involves at least 17 enzymes. It takes place primarily in the abundant smooth endoplasmic reticulum (SER).2 Steroidogenic cells have up to 10-fold more SER by volume than rough

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endoplasmic reticulum. Enzymes involved in steroid hormone formation and metabolism are also embedded in the SER. Trophic hormones that stimulate steroidogenesis generally increase both cellular cholesterol synthesis and lipoprotein uptake. It has been proposed that de novo cholesterol synthesis involves the production of several important local regulators of reproductive function in addition to substrate for steroid hormone synthesis. Biosynthetic intermediates between lanosterol and cholesterol have been shown to stimulate oocyte maturation in in vitro assays.11,12 These 4,4-dimethyl sterols, referred to as meiosis-activating sterols, contain 29 carbons and are found in the testis and follicular fluid in low micromolar concentrations. However, their physiological role in gamete maturation in vivo is uncertain based on results from germ cellspecific-knockout mouse models (targeted deletion of cytochrome P450 lanosterol 14α-demethylase, Cyp51a1) that indicate that de novo synthesis of meiosis-activating sterols is probably not essential for reproduction.13 The use of meiosis-activating sterols in clinical settings of animal and human reproduction remains a subject of debate, although there are reports of a positive impact on in vitro maturation of animal oocytes.

RELATIVE ROLES OF PLASMA LIPOPROTEINS AND DE NOVO SYNTHESIS IN SUPPLYING CHOLESTEROL FOR STEROIDOGENESIS The quantitative importance of circulating cholesterol carried by LDL, HDL, and other lipoproteins as a steroid hormone precursor in tissues that produce large amounts of hormone (e.g., the corpus luteum), as opposed to de novo cholesterol synthesis, is demonstrated by the fact that radiolabeled plasma cholesterol in humans is almost fully equilibrated with the steroidogenic pool of cholesterol. Inherited disorders have also shed light on the relative roles of lipoproteins and de novo cholesterol synthesis. Hypobetalipoproteinemia, a disorder in which there is virtually no circulating LDL,14,15 is a rare metabolic disorder associated with reduced adrenocortical steroid production and diminished progesterone levels during the luteal phase and in pregnancy. However, the lower levels of progesterone elaborated are still sufficient to support a term pregnancy. Hypercholesterolemia due to inactivating mutations in the LDL receptor gene is associated with modest impairment of steroidogenic gland function, reflecting the capacity of alternative sterol acquisition mechanisms to compensate for LDL receptor deficiency. In the case of HDL, individuals with a SR-B1 (SCARB1) missense variant (Pro297Ser) that reduces uptake

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function, have increased HDL levels and attenuated adrenal steroidogenesis in response to ACTH stimulation.10 SCARB1 polymorphisms associated low SR-B1 expression in granulosa cells have been reported to be accompanied by reduced estradiol levels and lower progesterone production. The commonly used cholesterol-lowering “statins,” which inhibit 3-hydroxy-3-methylglutaryl coenzyme A reductase (HMG-CoA reductase), the rate-limiting enzyme in de novo cholesterol synthesis, do not impair luteal steroidogenesis in adult humans despite the lowering of plasma LDL levels.16,17 Smith-Lemli-Opitz syndrome, an autosomal recessive disease, offers insight into the relationship of plasma cholesterol and de novo sterol synthesis for the supply of precursors for fetal steroidogenesis.18 The disease is caused by inactivating mutations in an enzyme involved in the terminal steps of cholesterol synthesis, 3β-hydroxysteroid Δ7-reductase (encoded by DHCR7). As a result, cholesterol levels are low and 7-dehydrocholesterol levels are elevated. Estrogen production during pregnancy is also reduced, due to impaired fetal adrenal hormone production. B-ring unsaturated equine-like steroids (1,3,5[10], 7-estratetrenes) are produced from the 7-dehydrocholesterol that accumulates, showing that the steroidogenic enzymes do not have an absolute requirement for cholesterol as a substrate.19 Desmosterolosis, another rare autosomal recessive disease caused by mutations in the 3β-hydroxysterol Δ24 reductase, is also associated with impaired fetal steroid synthesis.18

between lipid droplets and other organelles, including mitochondria. Mobilization of lipid droplet cholesterol esters occurs when cells are stimulated by trophic hormones. Adenosine 30 ,50 cyclic monophosphate (cAMP) triggers phosphorylation of perilipins by protein kinase A, and the subsequent detachment of perilipins from the droplet surface, allowing lipases access to the droplet sterol esters. The lipases that liberate free cholesterol from lipid droplet sterol esters are hormone-sensitive lipase and neutral cholesterol esterases (Fig. 2).6,23–25 Protein kinase A activates hormone sensitive lipase by phosphorylation of serine residues, promoting binding of the sterol esterase to lipid droplets. This enzyme’s role in steroidogenesis was suggested by reduced production of corticosterone under ACTH stimulation associated with an accumulation of lipid droplets in the adrenal cortex of mice deficient in Lipe. Targeted mutation of both the Lipe and Nceh1 genes in mice results in adrenal enlargement and lipid accumulation, but no impairment in

Mitochondrial membranes

Steroidogenic acute regulatory protein (STARD1)

Outer

START domain

Inner

Cholesterol Ser195 N-terminal mitochondrial targeting sequence

P450scc

LIPID DROPLETS Cytoplasmic lipid droplets represent a major depot of steroid hormone precursor. As much as 80% of the total cholesterol content of steroidogenic cells is esterified in the droplets. The sterol esters are synthesized in the endoplasmic reticulum from cholesterol acquired from lipoproteins or de novo synthesis by sterol Oacyltransferase 1 (SOAT1, previously named acylcoenzyme A, cholesterol acyltransferase-1 or ACAT1), encoded by one of two related genes.20 The esters generated by SOAT1 accumulate within the SER, and subsequently bud off as lipid droplets (Fig. 1). Targeted mutation of the Soat1 gene in mice results in markedly reduced sterol ester storage in the adrenal cortex without impairment of basal or ACTH-stimulated corticosterone production by adrenal cells. These findings suggest that transit through the sterol ester pool is not part of an obligatory itinerary for steroidogenic substrate. The limiting membranes of lipid droplets contain perilipins,21,22 proteins that protect the droplet contents from hydrolysis in the basal state. They also serve as scaffolds, anchoring lipases to the lipid droplet surface, as well as mediating physical and functional interactions

TOM

TIM

Steroidogenesis “ON” P450scc

TOM

TIM

Steroidogenesis “OFF” P450scc

FIG. 2 Structure of steroidogenic acute regulatory protein (STARD1) and model for its mechanism of action on intramitochondrial cholesterol translocation. StAR preprotein is folded by GRP78 in the endoplasmic reticulum and targeted to the outer mitochondrial membrane where it acts to promote cholesterol movement to the cholesterol side-chain cleavage enzyme (P450scc). StAR preprotein is subsequently imported by the translocator complex into the mitochondrial matrix where it is proteolytically processed to the mature 30 kDa form. TIM, inner mitochondrial membrane translocators; TOM, outer mitochondrial membrane protein translocators. Used with permission from Elsevier.5

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ACTH-stimulated corticosterone production. These different mouse models suggest a role for both hormonesensitive lipase and neutral cholesterol ester hydrolase activities in mobilization of sterol esters. The variable impact of enzyme deficiency on steroidogenesis in these murine models may be related to genetic background, or the age of mice at the time of study since lipid accumulation may have secondary deleterious effects on cell function beyond the disruption of sterol ester hydrolysis. The size and number of lipid droplets changes as the ester pool expands or contracts.26 The quantity of sterol ester stored is determined by the availability of cholesterol to the cell through de novo synthesis, through accumulation of lipoprotein-carried cholesterol, and by the steroidogenic activity of the cell. Trophic stimulation promotes cholesterol ester hydrolysis and diverts cholesterol into the steroidogenic pool away from SOAT1, preventing reesterification and resulting in a net depletion of cholesterol from the lipid droplets. Conversely, pharmacological blockade of steroid hormone synthesis (e.g., with the cholesterol side-chain cleavage inhibitor, aminoglutethimide) or defects in cholesterol use for steroidogenesis (e.g., congenital lipoid adrenal hyperplasia) increase sterol ester storage by increasing the amount of cholesterol available to SOAT1.2

INTRACELLULAR CHOLESTEROL TRAFFICKING The intracellular itineraries of lipoprotein-derived cholesterol, free cholesterol from the plasma membrane, or free cholesterol released from lipid droplets remains to be elucidated. In particular, much is still unknown about the ways in which sterol is presented to the mitochondria, where the first step in steroidogenesis takes place. The trafficking of cholesterol within ovarian cells is regulated by gonadotropins and impaired by endocrine disruptors like Bisphenol A. It is likely that sterol distribution to and from organelles occurs through a dynamic vesiculartubular late endosomal compartment, as well as through the assistance of lipid transfer proteins and proteins that establish contact sites between membranes that allow cholesterol to flow down a chemical gradient.2 The lipid transfer proteins involved in this process may include ATP-binding cassette transporter G1 (ABCG1), proteins with a structure resembling StAR (STARD1), including its paralogues, STARD3, STARD4, STARD5 (Fig. 4.2), and SCP2.2,7–9 The specific roles of these proteins remain to be clarified and their functions may be redundant since knockout mouse models with the exception of STARD1 do not show robust cholesterol metabolism/steroidogenic phenotypes. The mitochondria of steroidogenic cells are frequently found in close association with cytoplasmic lipid

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droplets, which may facilitate movement of substrate from these depots to the mitochondria through contact sites. Steroidogenic mitochondria have tubulovesicular cristae, in contrast to the lamellar cristae that are characteristic of mitochondria in other cells. The inner mitochondrial membranes contain the cholesterol side-chain cleavage enzyme, which catalyzes the first step in cholesterol metabolism into steroid hormones leading to the formation of pregnenolone. The hydrophobic cholesterol substrate must move from the mitochondrial outer membrane across the aqueous intermembranous space to reach the inner membrane. This translocation process is the rate-limiting step in steroidogenesis.2 The capacity to produce large amounts of steroid hormone in rapid response to trophic stimulation requires the action of STARD1, the prototypic member of the START domain family, which greatly enhances the flux of substrate to the side-chain cleavage system (Fig. 2).2 The mitochondrial cholesterol side-chain cleavage system is also juxtaposed to downstream enzymes in the steroidogenic pathway on the endoplasmic reticulum, allowing for efficient metabolism of pregnenolone.

REGULATION OF CELLULAR CHOLESTEROL BALANCE Cellular free cholesterol balance is highly regulated in cells by transcriptional and post-translational mechanisms. The expression of genes involved in cholesterol biosynthesis, like the gene encoding 3-hydroxy-3methylglutaryl coenzyme A reductase (HMGCR), and the uptake of LDL plasma cholesterol (LDLR) are controlled by master transcription factors, the sterol regulatory element-binding proteins (SREBF1, which encodes SREBP-1a and SREBPB-1c, and SREBF2, which encodes SREBP-2).2,27 SREBP-1a and SREBP-2 are the key transcription factors regulating cholesterol synthesis. The SREBPs are synthesized as inactive precursors that are bound to the endoplasmic reticulum. They interact with regulatory proteins, SREBP-cleavage activating protein (SCAP) and the insulin-induced genes 1 and 2, which inhibit SCAP. When cells are depleted of sterols, SCAP transports the SREBPs from the endoplasmic reticulum to the Golgi apparatus where SREBPs are cleaved through the action of two different proteases, releasing an NH2-terminal domain of the transcription factors that enter the nucleus where they activate genes controlling lipid synthesis and uptake. Cholesterol loading inhibits the movement of SREBPs to the Golgi and consequently the proteolytic processing, resulting in reduced transcription of the cholesterol synthesis and uptake genes. A micro ribonucleic acid RNA, miR-33, derived from an intron located within the gene encoding SREBP-2,

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posttranscriptionally modulates expression of genes involved in cellular cholesterol balance. Metabolites of cholesterol, the side-chain oxygenated sterols including 22-, 24-, 25-, and 27-hydroxycholesterol, and 7-hydroxylated sterols, are endogenous regulators of cellular cholesterol metabolism acting through the liver X receptors, LXRalpha and LXRbeta, also known as NR1H3 and NR1H2, respectively.28,29 In addition to LXRs, steroidogenic factor 1 (SF-1, also known as NR5A1) and a related protein liver receptor homolog-1 (LRH-1, also known as NR5A2) play roles in regulating genes encoding cholesterol metabolizing enzymes including SOAT1, LIPE, and STARD1. Other posttranslational mechanisms influencing sterol synthesis and uptake include cholesterol-induced ubiquitination of HMG-CoA reducatase, which tags the protein for degradation by proteasomes. Another posttranslational mechanism by which cells control sterol balance is the degradation of LDL receptors by proprotein convertase subtilisin/kexin 9 (PCSK9), a secreted serine protease that binds to LDL receptors on the cell surface and interferes with their recycling so that they are directed to lysosomes for degradation. Gain-of-function mutations in the PCSK9 gene cause autosomal dominant hypercholesterolemia, while loss-of-function mutation are associated with low LDL levels. Inhibition of PCSK9 reduces plasma LDL cholesterol in hypercholestrolemic subjects.30 Cells also control their sterol economy by reverse cholesterol transport (sterol efflux), mediated by members of the ATP-binding cassette subfamily A1 (ABCA1), which transfers cholesterol to plasma lipoproteins. In the primate corpus luteum undergoing functional luteolysis, a fall in STARD1 expression results in diminished progesterone production.31,32 Expression of lipoprotein receptors is reduced (reduced cholesterol uptake), and expression of ABCA1 is increased (increased cholesterol efflux), perhaps by hydroxysterol activation of LXR transcription factors. The excess cholesterol not used for steroidogenesis is esterified and deposited in cytoplasmic lipid droplets. These “homeostatic” adjustments reflect, in part, changes in gene transcription, posttranscriptional and post-translational processes that maintain optimal free cholesterol balance.

OVERVIEW OF STEROIDOGENESIS The cellular manufacture of steroid hormones involves the action of several classes of enzymes including the cytochromes P450, hemeprotein mixed-function oxidases named because of their distinct absorption peak at 450 nm when reduced in the presence of carbon monoxide, the hydroxysteroid dehydrogenases, and reductases.3

Cytochrome P450s catalyzes the major alterations in the sterol framework, cleavage of the side chain, hydroxylations, and aromatization. These hemeproteins require molecular oxygen and a source of reducing equivalents (i.e., electrons) to complete a catalytic cycle. Each member of the steroidogenic cytochrome P450 family of genes is designated “CYP,” followed by a unique identifying number that usually refers to the carbon atom at which the enzyme acts. The hydroxysteroid dehydrogenases reduce ketone groups or oxidize hydroxyl functions, employing pyridine nucleotide cofactors, usually with a stereospecific substrate preference and reaction direction. In addition to being involved in hormone biosynthesis in steroidogenic cells, this family of enzymes works with the reductases, steroid sulfotransferases, and steroid sulfatase to regulate the level of bioactive hormone in target tissues. The hydroxysteroid dehydrogenases are key determinants of the cellular response to endogenous steroid hormones as well as steroidal drugs. The reductases, using the pyridine nucleotide nicotinamide adenine dinucleotide phosphate (NADPH) as a cofactor, produce saturated ring A steroids from Δ4steroids (again, with stereospecificity). Here we will only discuss the enzymes involved in the initial steps in ovarian steroidogenesis.

KEY PROTEINS IN THE BIOSYNTHESIS AND METABOLISM OF STEROID HORMONES Steroidogenic Acute Regulatory Protein (STARD1): The Principal Regulator of Gonadal Steroidogenesis Translocation of cholesterol from the outer mitochondrial membranes to the relatively sterol-poor inner membranes is the critical step in steroidogenesis.2,3 This translocation process occurs at modest rates in the absence of specific effectors. It is markedly enhanced by STARD1, a protein with a short biological half-life. The following evidence established STARD1 as the key mediator of substrate flux to the cholesterol side-chain cleavage system: (1) Coexpression of STARD1 and the cholesterol side-chain cleavage enzyme system in cells that are not normally steroidogenic results in substantial pregnenolone synthesis above that produced by cells expressing the cholesterol side-chain cleavage enzyme system alone; (2) mutations that inactivate STARD1 cause congenital lipoid adrenal hyperplasia, a rare autosomal recessive disorder in which the synthesis of all adrenal and gonadal steroid hormones is severely impaired before the cholesterol side-chain cleavage step; (3) targeted deletion of the murine StarD1 gene results in a

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phenotype in nullizygous mice that mimics human congenital lipoid adrenal hyperplasia; and (4) STARD1 expression is directly correlated with steroidogenesis. Human STARD1 is synthesized as a 285-amino acid protein. The N-terminus of STARD1 is characteristic of proteins synthesized in the cytoplasm and then targeted and imported into mitochondria. Newly synthesized STARD1 preprotein (37 kDa) is rapidly imported into mitochondria and processed to the “mature” 30-kDa form by removal of the N-terminal amino acid sequence. The preprotein has a very short half-life (minutes), but the mature form is longer-lived (hours). The START domain consists of a unique structure that accommodates a cholesterol molecule. However, the binding of one sterol molecule is not sufficient to explain STARD1 action, and it is evident that STARD1 must be a catalyst for the transfer of multiple cholesterol molecules.2 STARD1 contains two consensus sequences for cyclic AMP (cAMP)-dependent protein kinase phosphorylation at Ser57 and Ser195. Ser195 of human STARD1 must be phosphorylated for maximal steroidogenic activity in model systems. Tissues that express STARD1 at high levels carry out trophic hormone-regulated mitochondrial sterol hydroxylations through the intermediacy of cAMP. STARD1 messenger RNA (mRNA) and protein are not present in the human placenta, an observation that is consistent with the fact that pregnancies hosting a fetus with mutations in the STARD1 gene causing congenital lipoid adrenal hyperplasia go to term. Although estrogen production is impaired in these pregnancies as a result of diminished fetal adrenal androgen production, placental progesterone synthesis is not significantly affected, indicating that the trophoblast cholesterol side-chain cleavage reaction is independent of STARD1. The abundance of STARD1 protein in steroidogenic cells is determined primarily by the rate of STARD1 gene transcription, which is influenced by transcription factors involved in the control of other genes involved in cholesterol metabolism including SREBPs, NR5A1, NR5A2, and LXRs (see below). STARD1 mRNA stability and translational mechanisms may also contribute. The STARD1 transcript is a novel target of microRNA let-7, which post-transcriptionally regulates STARD1 expression Let-7 is regulated by the long-noncoding RNA (lncRNA) H19, and overexpression of H19 in human and murine cells stimulates STARD1 expression by antagonizing let-7.33 In differentiated cells, the STARD1 gene is activated by the cAMP signal transduction cascade within 15–30 min. In differentiating cells (e.g., luteinizing granulosa cells), the induction of STARD1 transcription takes hours and requires ongoing protein synthesis. STARD1 was initially thought to stimulate cholesterol movement from the outer to the inner mitochondrial

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membrane as it was imported into the mitochondria. However, STARD1 protein lacking the mitochondrial targeting sequence is as effective as native STARD1 in stimulating steroidogenesis. Other STARD1 constructs engineered for prolonged tethering to the surface of the mitochondria were very active in stimulating pregnenolone production, suggesting that the residency time of the protein on the mitochondrial surface determines the duration of the steroidogenic stimulus. Recombinant human STARD1 lacking the N-terminal mitochondrial targeting sequence enhanced pregnenolone production by isolated ovarian mitochondria in a dose- and timedependent fashion, with significant increases in steroid production observed within minutes. Collectively, these findings strongly suggest that STARD1 acts on the outer mitochondrial membrane to promote cholesterol translocation. This perspective implies that import of the protein into the mitochondrial matrix, rather than being the trigger to steroid production, is actually the “off” mechanism, because it removes STARD1 from its site of action (Fig. 2). Consequently, on-going production of STARD1 preprotein is required to sustain steroidogenesis. The findings of the previously described experiments are most consistent with the idea that STARD1 enhances desorption of cholesterol from the sterol-rich outer mitochondrial membrane to the relatively sterol-poor inner membranes. The desorption process may involve a pH-dependent conformational change (molten globule transition). Even though STARD1 contains a hydrophobic pocket that binds cholesterol, sterol binding is not required for steroidogenic activity.2 Recent studies suggest that the endoplasmic reticulum chaperone, glucose regulatory protein 78 (GRP78), is associated with the mitochondrial membrane where it is thought to fold STARD1 for delivery to the outer mitochondrial membrane.34 The molecule or structure on the mitochondrial outer membrane that STARD1 acts on to promote cholesterol movement to the inner membrane has not been definitively identified. The STARD1 “receptor,” which could be a protein or a lipid, is not specific for mitochondria of steroidogenic cells, since STARD1 works in the context of COS-1 cells, which are not normally steroid hormone-producing cells. Thus, the specificity of the mechanism of mitochondrial cholesterol translocation is determined by whether STARD1 is expressed. Initially, the translocator protein (TSPO), also known as the peripheral benzodiazepine receptor, which is found in the outer mitochondrial membrane of many cell types, was postulated to be a target of STARD1 based on the fact that molecules that bind to TSPO stimulate steroidogenesis, and knockdown of TSPO expression in cultured cells reduced steroid synthesis in the presence of STARD1. However, the subsequent phenotyping of TSPO-deficient mice revealed that TSPO is dispensable

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for steroidogenesis. Despite on-going debate in the literature over the latter findings, the weight of evidence indicates that TSPO is not required for STARD1 action.35 Another potential candidate for the STARD1 effector molecule is the mitochondrial voltage-dependent anion channel 2 (VDAC2), which is also expressed by many different cell types, and has been proposed to be a cholesterol “pore” through which outer mitochondrial membrane cholesterol could flow to the inner membrane. As noted above, mutations in the STARD1 gene cause congenital lipoid adrenal hyperplasia, a rare autosomal recessive disease. Exceptions occur in Japan and Korea, however, where the mutation accounts for at least 5% of all cases of congenital adrenal hyperplasia.2,36 The pathophysiology of the disease entails a two-step process in which impaired use of cholesterol for steroidogenesis leads to accumulation of sterol esters in lipid droplets. These droplets ultimately compress cellular organelles, causing damage through the formation of lipid peroxides. Mutations inactivating NR5A1 (SF-1), a master transcription factor controlling expression of steroidogenic genes, also cause cholesterol accumulation because STARD1 as well as cholesterol side-chain cleavage enzyme expression are impaired. Mutations found in the STARD1 gene, which is composed of seven exons, include frameshifts caused by deletions or insertions, splicing errors, and nonsense and missense mutations. All of these mutations lead to the absence of STARD1 protein or the production of functionally inactive protein. Several nonsense mutations were shown to result in C-terminus truncations of STARD1. One of these mutations, Gln258Stop, results in the deletion of the final C-terminal 28 amino acids of the STARD1 protein and accounts for 80% of the known mutant alleles in the affected Japanese population. Known point mutations that produce amino acid substitutions occur in exons 5–7 of the gene, the exons that encode the C-terminus. Mutations that cause partial loss of STARD1 activity (usually 20%–30% of normal) are associated with a milder disease phenotype or “nonclassical” disease.2 Although affected XY subjects are pseudohermaphrodites (46,XY disorder of sexual development (DSD)) because of an inability to generate sufficient fetal testicular testosterone to masculinize the external genitalia, XX subjects have normal external genitalia, develop female secondary sexual characteristics, and experience menarche. They are, however, anovulatory and unable to produce large amounts of estradiol and progesterone in a cyclic fashion. The fact that some ovarian estradiol synthesis occurs reflects the existence of STARD1independent substrate movement to the cholesterol side-chain cleavage system. Pregnancy through assisted reproduction technology has been achieved in a XX female with congenital lipod adrenal hyperplasia and no spontaneous puberty after ovarian stimulation,

in vitro fertilization and transfer of frozen-thawed embryos after endometrial preparation with exogenous hormones.37

The Cholesterol Side-Chain Cleavage Enzyme (P450scc Encoded by CYP11A1) Cholesterol side-chain cleavage is catalyzed by cytochrome P450scc and its associated electron transport system, consisting of a flavoprotein reductase (ferredoxin or adrenodoxin reductase) and an iron sulfoprotein (ferredoxin or adrenodoxin), encoded by the FDX1 gene, which shuttles electrons to cytochrome P450scc.2,3,38,39 The sidechain cleavage reaction involves three catalytic cycles: the first two lead to the introduction of hydroxyl groups at positions C-22 and C-20, and the third results in scission of the side chain between these carbons. Each catalytic cycle requires one molecule of NADPH and one molecule of oxygen so that the formation of one mole of the cleavage products, pregnenolone and isocroapraldehyde, uses three moles of NADPH and three moles of oxygen. The slowest step of the reaction is the binding of cholesterol to the hydrophobic pocket of P450scc, where the heme resides. The sterol substrate remains bound to a single active site on cytochrome P450scc for all three cycles because of the tight binding of the reaction intermediates. The dissociation constant (Kd) for binding of cholesterol, a measure of the enzyme’s affinity for its substrate, is approximately 5000 nM, whereas the Kd for the binding of the intermediate product 22-hydroxycholesterol is 4.9 nM; the Kd for 20,22-dihydroxycholesterol is 81 nM. The estimated Kd for pregnenolone, the end product, is 2900 nM, which permits its dissociation from the enzyme at the end of the reaction. Reducing equivalents are shuttled to cytochrome P450scc by ferredoxin in cycles of reduction and oxidation, facilitated by differential affinities of the proteins, depending on their state of oxidation or reduction.39 Ferredoxin forms a 1:1 complex with ferredoxin reductase, which catalyzes reduction of the iron-sulfur protein. The reduced ferredoxin then dissociates and forms a 1:1 complex with cytochrome P450scc and is subsequently oxidized when it donates its electrons to P450scc. Oxidized ferredoxin returns to ferredoxin reductase for electron recharging. This recharging is facilitated by the fact that ferredoxin reductase has a greater affinity for oxidized over reduced ferredoxin. The binding of cholesterol to cytochrome P450scc increases its affinity for reduced ferredoxin, which enhances the shuttle of electrons to substrate-loaded enzyme. The rate of formation of pregnenolone is determined by: (1) Access of cholesterol to the inner mitochondrial membranes; (2) the quantity of cholesterol side-chain cleavage enzyme, and secondarily, its flavoprotein and

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iron-sulfur protein electron transport chain; and (3) catalytic activity of P450scc, which can be influenced by post-translational modification. Acute alterations in steroidogenesis generally result from changes in the delivery of cholesterol to P450scc, whereas long-term alterations involve changes in the quantity of enzyme proteins as well as cholesterol delivery.2,3 Mutations in the CYP11A1 gene that result in significantly diminished cholesterol side-chain cleavage activity have been reported in association with adrenal insufficiency and XY DSD (sex reversal), phenotypes that are similar to those associated with inactivating mutations in the STARD1 gene.2,40–42 One interesting CYP11A1 mutation was discovered in an XY subject, born prematurely with sex reversal and adrenal failure. The subject was homozygous for a single-nucleotide deletion leading to a premature termination codon at codon 288, predicted to delete the C-terminal 242 amino acids, including the heme-binding site, which should result in a nonfunctional protein. However, other subjects with inactivating CYP11A1 mutations are born at term and have later onset of adrenal insufficiency. Knockout of the Cyp11a1 gene in mice produces a generally similar phenotype. The discovery of mutations causing severe P450scc deficiency in some humans born at term challenges the notion that absence of P450scc activity in the fetus and placenta is incompatible with pregnancy progressing beyond the usually limited period of luteal progesterone support. Therefore, compensatory mechanisms appear to exist to maintain sufficient progestational activity, perhaps sustained corpus luteum function, to sustain pregnancy and fetal viability in women hosting a fetus with CYP11A1 mutations.

3β-Hydroxysteroid Dehydrogenase/Δ5-4 Isomerase The 3β-HSD/Δ5-4 isomerases are membrane-bound enzymes localized to the endoplasmic reticulum and mitochondria that use nicotinamide adenine dinucleotide (NAD +) as a cofactor.43 These enzymes catalyze dehydrogenation of the 3β-hydroxyl group and the subsequent isomerization of the Δ5 olefinic bond to yield a Δ4 ketone structure. They convert pregnenolone into progesterone, 3β-17α-hydroxypregnenolone into 17α-hydroxyprogesterone, and dehydroepiandrosterone into androstenedione.3,43 The dehydrogenase and isomerase reactions are performed at a single bifunctional catalytic site that adopts different conformations for each activity. The 3β-hydroxysteroid dehydrogenase step is rate limiting in the overall reaction sequence, and the NADH formed in this reaction is believed to alter the enzyme conformation to promote the isomerase reaction.

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The human genome has two active 3β-HSD/Δ5-4 isomerase genes: one encodes a protein predominantly expressed in the placenta, liver, breast, and brain (HSD3B1), the other a protein expressed in the gonads and adrenal cortex (HSD3B2). These two genes, each consisting of four exons each, lie 100 kb apart on band 1p13.1. The human genome also contains five unprocessed pseudogenes closely related to HSD3B1 and HSD3B2 on band 1p13.1, with two of them lying between the expressed genes. The DNA sequences of the exons of the two active genes are very similar, and the encoded proteins differ in only 23 amino acid residues. HSD3B1, however, has a lower Km for substrate than HSD3B2), which facilitates metabolism of lower concentrations of Δ5 substrate. Electron microscope cytochemistry has localized HSD3B2 activity to the perimitochondrial endoplasmic reticulum and in subcellular fractions containing STARD1 and P450scc. In some cell types, the enzymes appear to be localized in the inner mitochondrial membrane. Thus, they are positioned to act on pregnenolone produced by the cholesterol side-chain cleavage system. Because most steroidogenic cells have a large capacity to generate progesterone when presented with exogenous pregnenolone, the 3β-HSD/Δ5-4 isomerases are not believed to be rate-determining enzymes. However, mutations causing deficiency of HSD3B2 cause a form of congenital adrenal hyperplasia characterized by impaired gonadal and adrenal steroidogenesis with accumulation of Δ5 steroids in the circulation. In its severest form, HSD3B2 deficiency is associated with salt wasting because of insufficient mineralocorticoid production.3,43,44 Kinetic analysis of mutant proteins associated with the salt-wasting and non-saltwasting forms of the disease showed a four to fortyfold reduction in catalytic efficiency for the conversion of pregnenolone into progesterone. The salt-wasting form of the disease is associated with frameshift mutations resulting in protein truncation and a variety of missense mutations that affect affinity for the cofactor and protein stability. The greater instability of the mutant proteins found in subjects with salt-wasting disease compared with those proteins found in the non-salt-wasting form appears to account, in part, for the different clinical phenotypes. A so-called attenuated, or late-onset, form of 3β-HSD deficiency, diagnosed by steroid measurements, has been described in the literature. However, no mutations have yet been discovered in the genes encoding HSD3B1 and HSD3B2 in subjects with this clinical diagnosis. Mutations in the distal promoter or epigenetic factors that might alter enzyme expression cannot be ruled out. The apparent reduced 3β-HSD activity could also be the result of alterations in the membrane environment that affect catalytic activity or posttranslational modifications to the enzyme that diminish its function.

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REGULATION OF EXPRESSION OF THE STEROIDOGENIC MACHINERY The expression of genes encoding proteins involved in cholesterol acquisition and its transformation into steroid hormones is controlled by gonadotropins, whose actions are modulated by locally acting paracrine and autocrine factors within the ovary (e.g., GDF-9 derived from the oocyte and granulosa cells inhibits granulosa cell and thecal steroidogenesis). Steroidogenesis is subject to both long-term control through the process of differentiation, for example, the differentiation of granulosa cells into granulosa lutein cells on ovulation, which is associated with a huge increase in the capacity to synthesize progesterone; and acute regulation, primarily driven by LH, as seen in the late luteal ophase spikes of progesterone production by the corpus luteum in response to pulsatile LH release. The regulation of expression of genes encoding proteins involved in steroidogenesis in the ovary, testes, and adrenal cortex shares a number of similarities with respect to the involvement of cis elements and transcription factors. SF-1, a nuclear receptor, also known as Ad4BP and by the nuclear receptor family designation NR5A1, is essential for development of steroidogenic glands.45,46 Most of the genes encoding key proteins involved in steroidogenesis (e.g., SCARB1, STARD1, CYP11A1, CYP11B2, CYP17A1, CYP19A1, CYP21A2) contain one or more NR5A1 response elements in their proximal promoters. These elements are important for basal as well as stimulated expression of these genes, generally by a cAMP-mediated signal transduction pathway. The importance of cAMP signaling in steroidogenic tissues is reflected in increased steroid production when specific phosphodiesterases are blocked, which elevates intracellular cAMP levels, or when mutations occur in the gene encoding the regulatory subunit of protein kinase A (PRKAR1A), resulting in unrestrained protein kinase A activity.3 There appear to be several mechanisms by which NR5A1 action is modified by cAMP action, including phosphorylation of the protein by kinases, providing a link between this transcription factor and molecules that transduce signals from plasma membrane receptors.47 The importance of NR5A1 to the regulation of steroidogenic tissues was documented by gene targeting. Mice deficient in NR5A1 lacked adrenal glands and gonads, and males were consequently sex reversed. Haploinsufficiency of Nr5α1 in the mouse resulted in an impaired adrenal steroidogenic response to stress, although basal steroidogenesis was not affected due to compensatory hypertrophy. NR5A1 haploinsufficiency has been reported in humans with primary adrenal failure and XY DSD. It is now evident that variation in the NR5A1 gene is associated with a range of phenotypes

such as 46,XYDSD, hypospadias, anorchia, male factor infertility, and primary ovarian insufficiency. Among the NR5A1 mutations reported are missense mutations within the DNA-binding region, a nonsense mutation, a and a frameshift mutation predicted to disrupt RNA stability or protein function. Functional studies of the missense mutants (Cys33Ser, Arg84His) and of one nonsense mutant (Tyr138Stop) showed impaired activation of NR5A1-responsive target genes. In addition to NR5A1, other transcription factors participate in the control of genes encoding the steroidogenic machinery. A related transcription factor, liver receptor homolog-1 (SF-2 or NR5A2) recognizes the same canonical DNA motif to which NR5A1 binds and may share functions with NR5A1 in certain tissues, including the adrenal cortex, testis, and ovary.48 Both NR5A1 and NR5A2 have been crystallized and found to contain phospholipid-binding pockets, with phosphatidyl inositols being the presumed ligands. These observations suggest that phospholipids may be regulatory molecules controlling expression of genes involved in steroidogenesis.49 The tissue-specific regulation of genes expressed in multiple steroidogenic glands (e.g., CYP17A1) requires the action of other transcription factors working either independently or in concert with NR5A1 in a combinatorial fashion. In addition, the activity of NR5A1 is regulated by transcription factors that either bind to NR5A1 response elements and prevent activation of transcription (chicken ovalbumin upstream promoter-transcription factor (COUP-TF)) or bind to NR5A1 and block its ability to transactivate promoters (DAX-1, also known as NR0B1). Interestingly, expression of the latter gene is upregulated by NR5A1, so there is a complex control mechanism in place for modulating these antagonistic molecules. Other transcription factors that are known to be important for the expression of genes involved in steroidogenesis include GATA4 and GATA6, members of the GATA family of transcription factors originally identified as being central to hematopoiesis and endoderm development, and LXRalpha.50,51

CONCLUSION Ovarian steroidogenic cells are primarily controlled by pituitary gonadotropins that promote cellular differentiation, leading to increased capacity to produce steroid hormones, and to the acute stimulation of steroidogenesis in the differentiated state, primarily through the intermediacy of steroidogenic acute regulatory protein, which mediates the transfer of cholesterol from the outer mitochondrial to the inner mitochondrial membrane where

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REFERENCES

the first biosynthetic step in steroid synthesis, cholesterol side-chain cleavage, occurs. Each steroidogenic cell type of the ovary has unique attributes, but they are all endowed with the ability to acquire cholesterol from lipoprotein particles or it synthesize de novo. An overlapping repertoire of transcription factors coordinates the expression of the enzymes and proteins required for the process of hormone synthesis.

References [1] Hu J, Zhang Z, Shen WJ, Azhar S. Cellular cholesterol delivery, intracellular processing and utilization for biosynthesis of steroid hormones. Nutr Metab (Lond) 2010;1(7):47. [2] Miller WL. Disorders in the initial steps of steroid hormone synthesis. J Steroid Biochem Mol Bol 2017;165:18–37. [3] Miller WL, Auchus RJ. The molecular biology, biochemistry, and physiology of human steroidogenesis and its disorders. Endocr Rev 2011;32(1):81–151. [4] Conley AJ, Corbin CJ, Thomas JL, et al. Costs and consequences of cellular compartmentalization and substrate competition among human enzymes involved in androgen and estrogen synthesis. Biol Reprod 2012;86(1):1–8. [5] Strauss JF, FitzGerald GA. Steroid hormones and other lipid molecules involved in human reproduction. In: Strauss JF, Barbieri RL, editors. Yen and Jaffe’s reproductive endocrinology: physiology, pathophysiology, and clinical management. 8th ed. Philadelphia: Elsevier; 2019. [6] Kraemer FB. Adrenal cholesterol utilization. Mol Cell Endocrinol 2007;265–266:42–5. [7] Maxfield FR, Iaea DB, Pipalla NH. Role of STARD4 and NPC1 in intracellular sterol transport. Biochem Cell Biol 2016;94:499–506. [8] Li NC, Fan J, Papadopoulos V. Sterol carrier protein-2, a nonspecific lipid-transfer protein, in intracellular cholesterol trafficking in testicular Leydig cells. PLoS One 2016;11(2):e0149728 https:// doi.org/10.1371/journal.pone.0149728. [9] Wilhelm LP, Tomasetto C, Alpy F. Touche! STARD3 and STARD3NL tether the ER to endosomes. Biochem Soc Trans 2016;44(2):493–8. https://doi.org/10.1042/BST20150269. [10] Hoekstra M, Van Eck M, Korporaal SJ. Genetic studies in mice and humans reveal new physiological roles for the high-density lipoprotein receptor scavenger receptor class B type I. Curr Opin Lipidol 2012;23:127–33. [11] Grøndahl C. Oocyte maturation. Basic and clinical aspects of in vitro maturation (IVM) with special emphasis of the role of FF-MAS. Dan Med Bull 2008 Feb;55(1):1–16. [12] Keber R, Rozman D, Horvat S. Sterols in spermatogenesis and sperm maturation. J Lipid Res 2013;54(1):20–33. [13] Keber R, Acimovic J, Majdic G, Motaln H, Rozman D, Horvat S. Male germ cell-specific knockout of cholesterogenic cytochrome P450 lanosterol 14α-demethylase (Cyp51). J Lipid Res 2013;54 (6):1653–61. [14] Illingworth DR, Corbin DK, Kemp ED, et al. Hormone changes during the menstrual cycle in abetalipoproteinemia: reduced luteal phase progesterone in a patient with homozygous hypobetalipoproteinemia. Proc Natl Acad Sci U S A 1982;79:6685. [15] Parker Jr. C, Illingworth DR, Bissonnette J, et al. Endocrine changes during pregnancy in a patient with homozygous familial hypobetalipoproteinemia. N Engl J Med 1986;314:557. [16] Plotkin D, Miller S, Nakajima S, et al. Lowering low density lipoprotein cholesterol with simvastatin, a hydroxy-3-metyhylglutarylcoenzyme A reductase inhibitor, does not affect luteal function in premenopausal women. J Clin Endocrinol Metab 2002;87:3155.

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[17] Laue L, Hoeg J, Barnes K, et al. The effect of mevinolin on steroidogenesis in patients with defects in the low density lipoprotein receptor pathway. J Clin Endocrinol Metab 1987;64:531. [18] Porter FD, Herman GE. Malformation syndromes caused by disorders of cholesterol synthesis. J Lipid Res 2011;52(1):6–34. [19] Shackleton CH, Roitman E, Kratz LE, et al. Equine type estrogens produced by a pregnant woman carrying a Smith-Lemli-Opitz syndrome fetus. J Clin Endocrinol Metab 1995;84:1157. [20] Chang TY, Chang CCY, Chen D. Acyl-coenzyme A: cholesterol acyltransferase. Annu Rev Biochem 1997;66:613. [21] Brasaemle DL. The perilipin family of structural lipid droplet proteins: stabilization of lipid droplets and control of lipolysis. J Lipid Res 2007;48:2547. [22] Wang H, Sreenevasan U, Hu H, et al. Perilipin 5, a lipid dropletassociated protein, provides physical and metabolic linkage to mitochondria. J Lipid Res 2011;52(12):2159–68. [23] Ohta K, Sekiya M, Uozaki H, et al. Abrogation of neutral cholesterol ester hydrolytic activity causes adrenal enlargement. Biochem Biophys Res Commun 2011;404(1):254–60. [24] Kraemer F, Shen W, Natu V, et al. Adrenal neutral cholesteryl ester hydrolase: identification, subcellular distribution, and sex difference. Endocrinology 2002;143:801. [25] Li H, Brochu M, Wang SP, et al. Hormone-sensitive lipase deficiency in mice causes lipid storage in the adrenal cortex and impaired corticosterone response to corticotropin stimulation. Endocrinology 2002;143:3333. [26] Strauss III JF, Schuler LA, Rosenblum MF, et al. Cholesterol metabolism by ovarian tissue. Adv Lipid Res 1981;18:99. [27] Ye J, DeBose-Boyd RA. Regulation of cholesterol and fatty acid synthesis. Cold Spring Harb Perspect Biol 2011;3(7). pii: a004754. [28] Brown AJ, Jessup W. Oxysterols: sources, cellular storage and metabolism, and new insights into their roles in cholesterol homeostasis. Mol Aspects Med 2009;30(3):111–22. [29] Cummins CL, Volle DH, Zhang Y, et al. Liver X receptors regulate adrenal cholesterol balance. J Clin Invest 2006;116(7):1902–12. [30] Tavori H, Rashid S, Fazio S. On the function and homeostasis of PCSK9: reciprocal interaction with LDLR and additional lipid effects. Atherosclerosis 2015;238(2):264–70. [31] Devoto L, Fuentes A, Kohen P, et al. The human corpus luteum: life cycle and function in natural cycles. Fertil Steril 2009;92 (3):1067–79. [32] Bogan RL, Hennebold JD. The reverse cholesterol transport system as a potential mediator of luteolysis in the primate corpus luteum. Reproduction 2010;139(1):163–76. [33] Men Y, Fan Y, Shen Y, Lu L, Kallen AN. The sterioidogenic acute regulatory protein (StAR) is regulated by the H19/let-7 axis. Endocrinology 2017;158:402–9. [34] Prasad M, Pawlak KJ, Burak WE, Perry EE, Marshall B, Whittal RM, Bose HS. Mitochondrial metabolic regulation by GRP78. Sci Adv 2017;33:e1602038. [35] Selvaraj V, Stocco DM, Tu LN. Translocator protein (TSPO) and steroidogenesis: a reappraisal. Mol Endocrinol 2015;29(4):490–501. [36] Bose HS, Sugawara T, Strauss III JF, et al. The pathophysiology and genetics of congenital lipoid adrenal hyperplasia. N Engl J Med 1996;335:1870. [37] Albarel F, Perrin J, Jegaden M, Roucher-Boulez F, Reynaud R, Brue T, Courbiere B. Successful IVF pregnancy despite inadequate ovarian steroidogenesis due to congenital lipoid adrenal hyperplasia (CLAH): a case report. Hum Reprod 2016;31(11):2609–12. [38] Heyl BL, Tyrell DJ, Lambeth JD. Cytochrome P-450scc-substrate interactions: roles of the 3-band side-chain hydroxyls in binding to oxidized and reduced forms of the enzyme. J Biol Chem 1986;261:2743. [39] Lambeth JD, Seybert DW, Kamin H. Phospholipid vesiclereconstituted cytochrome P-450scc: mutually facilitated binding of cholesterol and adrenodoxin. J Biol Chem 1980;255:138.

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[40] Katsumata N, Ohtake M, Hojo T, et al. Compound heterozygous mutations in the cholesterol side-chain cleavage enzyme gene (CYP11A) cause congenital adrenal insufficiency in humans. J Clin Endocrinol Metab 2002;87:3808. [41] Hiort O, Holterhus PM, Werner R, et al. Homozygous disruption of P450 side-chain cleavage (CYP11A1) is associated with prematurity, complete 46, XY sex reversal, and severe adrenal failure. J Clin Endocrinol Metab 2005;90:538. [42] Chien Y, Rosal K, Chung B-C. Function of CYP11A1 in the mitochondria. Mol Cell Endocrinol 2017;441:55–61. [43] Simard J, Ricketts M-L, Gingras S, et al. Molecular biology of the 3ß-hydroxysteroid dehydrogenase/Δ5-Δ4 isomerase gene family. Endocr Rev 2005;226:525. [44] Krone N, Arlt W. Genetics of congenital adrenal hyperplasia. Best Pract Res Endocrinol Metab 2009;23(2):181–92. [45] Schimmer BP, White PC. Minireview: steroidogenic factor 1: its roles in differentiation, development, and disease. Mol Endocrinol 2010;24(7):1322–37. [46] Ferraz-de-Souza B, Lin L, Achermann JC. Steroidogenic factor-1 (SF-1, NR5A1) and human disease. Mol Cell Endocrinol 2011;336 (1–2):198–205.

[47] Auchus RJ. The backdoor pathway to dihydrotestosterone. Trends Endocrinol Metab 2004;15(9):432–8. [48] Fayard E, Auwerx J, Schoonjans K. LRH-1: an orphan nuclear receptor involved in development, metabolism and steroidogenesis. Trends Cell Biol 2004;14:250. [49] Krylova IN, Sablin EP, Moore J, et al. Structural analyses reveal phosphatidyl inositols as ligands for the NR5 orphan receptors SF-1 and LRH-1. Cell 2005;120:343. [50] Martin LJ, Taniguchi H, Robert NM, et al. GATA factors and the nuclear receptors, steroidogenic factor 1/liver receptor homolog 1, are key mutual partners in the regulation of the human 3βhydroxysteroid dehydrogenase type 2 promoter. Mol Endocrinol 2005;19:2358–70. [51] King SR, Lavoie HA. Gonadal transactivation of STARD1, CYP11A1 and HSD3B. Front Biosci 2012;17(1):824–46.

Further Reading [52] Luo J, Jiang L, Yang H, Song B-I. Routes and mechanisms of postendosomal cholesterol trafficking: a story that never ends. Traffic 2017;12471: doi: 101111/tra.

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6 Inhibin, Activin, and Follistatin in Ovarian Physiology Corrine Welt, Alan Schneyer BIOCHEMISTRY AND MOLECULAR BIOLOGY OF INHIBIN, ACTIVIN, AND FOLLISTATIN

was produced not by sequencing the entire protein, which would have required large quantities of pure inhibin, but from its cDNA [8]. This combination of protein purification and molecular sequencing identified several features of inhibin structure. First, inhibin was composed of two disulfide-linked subunits designated alpha and beta. Moreover, inhibin was produced from larger precursor proteins that were proteolytically cleaved to form mature, 32,000 MW inhibin whose sequence and processing were identified from the nucleotide sequence. In this regard, the α-subunit was distinct from the β-subunit in that it had two cleavage sites creating pre-pro, pro, and mature fragments, which produced a stand-alone version of the α-subunit called pro-α-C as well as the mature dimerization partner for β-subunits [9]. Additional β-subunits were identified and named βB, βC, βD, and βE over the years. βA and βB could be linked to α-subunit to form inhibin A and inhibin B, respectively [9]. Finally, the structure and sequence was found to be similar to that of TGFβ, which had been purified and sequenced earlier [8]. This put inhibin within the enlarging TGFβ superfamily that now contains 30–50 members depending on how strictly the families are defined. Purification of inhibin using the pituitary cell bioassay had some additional advantages. Once the inhibin structure was defined, side fractions from the original purification were reanalyzed. A factor that specifically stimulated FSH release from gonadotropes was identified, named activin, which turned out to be a homodimer of β-subunits, creating activin A (2 βA subunits), activin B (2 βB subunits), and activin AB with one of each [10]. Another side fraction had weaker FSH inhibitory activity that was eventually called follistatin. Follistatin consisted of a single protein chain of approximately 35 kD [11] that was totally unrelated to inhibin or activin. The function of this protein was somewhat mysterious given that a more

In 1923, Mottram and Cramer [1] reported that irradiation of rat testes damaged seminiferous tubules. Interestingly, this treatment also caused the histology of pituitary cells to be radically altered by the appearance of “castration cells” or cells that have secreted the majority of their storage vesicles, suggesting that a factor regulating these pituitary cells comes from testes and is lost due to castration or irradiation. Shortly thereafter, McCullagh [2] reported that an aqueous testicular extract devoid of steroid hormones was sufficient to prevent formation of castration cells, thereby providing evidence for a proteinaceous substance that was secreted from the testis, traveled through the blood and regulated pituitary cell function, that is, an endocrine hormone. He coined the name “inhibin” for this biological activity. This observation set off a 60-year international effort to purify and identify the molecular nature of inhibin. Further advances required identification of the pituitary gonadotrope hormones that regulate gonadal function in males and females, follicle-stimulating hormone (FSH) and luteinizing hormone (LH), and development of assays to measure FSH and LH. With these developments, it became clear that the near complete release of FSH from gonadotropes after gonadectomy produced the “castration cell” appearance. This critical observation provided a reliable bioassay, first in vivo and later, using pituitary cells in culture, that could be used to purify inhibin from gonadal extracts based on its ability to specifically inhibit FSH release without affecting LH [3]. As it turns out, authentic inhibin was finally purified by four different groups in 1985 [4–7] at the time when molecular biology was becoming more widely applied in endocrinology. Thus, the first complete sequence for inhibin

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active gonadal FSH release inhibitor (i.e., inhibin) was produced by the same tissues. However, follistatin was determined to inhibit FSH by binding and neutralizing activin [12]. Thus, activin stimulated FSH production and release while follistatin inhibited this action by binding and neutralizing activin, establishing follistatin as a paracrine/autocrine regulator of this new feedback system. The final protein in this story is follistatin like-3 (FSTL3, aka follistatin-related gene; follistatin-related protein), which was discovered in the mid-1990s based on sequence similarity to follistatin [13]. FSTL3 is structurally very close to follistatin suggesting that they originated from a common ancestor, but have different production profiles and activities [14]. With respect to ovarian physiology, FSTL3 does not appear to have important actions [15] and will therefore not be discussed further. The last decade of the 20th century and the first of the 21st saw a virtual explosion of biochemical, molecular, and physiological studies defining important details of production and action of inhibin, activin, and follistatin in regulating reproduction in males and females. In addition, development of assays, first a radioimmunoassay [16] followed by more specific two-site ELISA assays, allowed investigation of inhibin, activin, and eventually follistatin in animal and human fluids [17]. Most of this work has been compiled and reviewed in a number of scholarly compilations (e.g., Ref. [18]). Moreover, most of the recent advances in this field have been in translation of the basic findings to clinical observations and applications. Therefore, we aim to provide an update from the earlier work that is the basis for current and future clinical applications. It should also be kept in mind that these proteins have actions outside of reproduction that might also be clinically important but won’t be covered here.

STRUCTURAL FEATURES, SIGNALING, AND REGULATION OF INHIBIN, ACTIVIN, AND FOLLISTATIN ACTIVITY As mentioned above, both inhibin α-subunit, and the inhibin/activin β-subunits are produced as pro-protein precursors that are cleaved by furin proteases to produce mature, dimeric proteins [9]. While the activins are inactive until the propeptide is cleaved, bioactive inhibins of various sizes have been identified containing unprocessed or partially processed α- or β-subunits in the inhibin dimer [19]. Whether these inhibin isoforms have different activities or just collectively comprise the “inhibin” signal remains to be determined. However, the most N-terminal fragment of the α-subunit precursor is secreted separately from mature inhibin and was recently found to regulate inhibin activity by binding to

the site where mature α-subunit binds to the betaglycan inhibin receptor [20] although the physiological significance of this observation is unknown. Since cleavage of the propeptide from activin is required for its activity it has been assumed that this portion of the molecule is required only for proper dimerization within the endoplasmic reticulum. However, other TGFβ superfamily members are secreted as complexes of the mature and propeptides that are cleaved but still inactive. An example is TGFβ itself, which is activated by proteolytic release from the propeptide at its site of action [21]. The activin propeptide was found to associate with mature activin but the affinity of this interaction was too low to neutralize activin actions as in the TGFβ situation [22]. By engineering a propeptide sequence that matched TGFβ in the C-terminal region but matched activin A in the N-terminal region, the affinity of this propeptide was vastly increased to the point that it became a specific antagonist for activins A or B [23] although still less potent but more specific for activin than follistatin. This propeptide can therefore be used to specifically inhibit activin activity in vivo [24]. In addition to sharing an overall molecular structure, members of the TGFβ superfamily also share a family of receptors for transducing their signals and a family of second messengers for transmitting signals to subcellular compartments including the nucleus. Activins signal through a heterotetrameric complex comprised of one of two ligand-binding type 2 receptors, ActRIIA or B (ACVR2A or B, respectively) that induce a conformational change in activin that opens up binding sites for the signal-inducing ActR1B (Alk4 or ACVR1B) receptors (reviewed in Ref. [18]). The inhibin receptor consists of ActRIIA or B in association with the betaglycan co-receptor with contacts to ActRII being conducted by the inhibin β-subunit while contacts with betaglycan are mediated by the α-subunit [25,26]. Thus, inhibin can act, at least in part, by binding to and therefore sequestering type II activin receptors that are then no longer available for activin signaling. Follistatin and FSTL3 both inhibit activins A and B [14,27,28]) by forming a complex of two follistatin molecules arranged head to tail around the activin dimer [29,30], an arrangement that covers up receptor binding regions on the ligand as well as creating a stable complex with a very slow off rate [27]. Therefore, both follistatin and FSTL3 are relatively high affinity antagonists of activins A and B. Once activin associates with its receptor complex a conformational change is induced that results in phosphorylation of the ActR1B receptor, which then phosphorylates the second messengers Smad2 or Smad3 [31]. These same Smads are also used by TGFβ itself acting via TGFBR1(ALK5) and it remains unknown how these signals are differentiated at the DNA level from activin-stimulated signaling. Nevertheless, activated

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Smad2 or 3 binds with a common Smad4 and the complex then migrates to the nucleus where it induces alterations in gene transcription.

SYNTHESIS AND ACTION OF INHIBIN, ACTIVIN, AND FOLLISTATIN IN THE OVARY The primary source of inhibin is gonadal since it rapidly becomes undetectable after castration [32]. Activins, although also produced in gonads, are produced in many other tissues so that the source(s) of circulating activin remains enigmatic. Moreover, activin may be more relevant as an autocrine/paracrine modulator of cellular physiology rather than an endocrine hormone, which has made elucidating its biological actions more challenging [31]. Similarly, serum levels of follistatin are unaffected by gonadal status [33] suggesting that locally produced follistatin within the ovary may have more critical actions regulating activin-induced ovarian development and follicular maturation. The primary action of follistatin remains binding and neutralizing activin and related TGFβ family ligands [34] so that the role of activin and follistatin in ovarian development and physiology is tightly intertwined [18]. Although FSH has been known to stimulate secretion of inhibin A and B from ovarian granulosa cells, the differential regulation of inhibin A and B during the human menstrual cycle, taken in conjunction with FSH secretory patterns suggests that this regulation is more complicated (reviewed in Ref. [18]). Inhibin B increases during the luteal-follicular transition as one menstrual cycle ends and the subsequent one begins. It rises to a peak in the mid-follicular phase, with a second peak on the day after the LH surge. Inhibin A on the other hand begins to rise late in the follicular phase reaching one peak at mid-cycle and another during the mid-luteal phase. This suggests that regulation of inhibin A and B production and secretion is not identical nor is their activity at the pituitary [18]. While purified inhibin A and B both suppress FSH in pituitary cell bioassays in vitro [9] the relative roles of inhibins A and B have yet to be established in the human. Further, inhibin overlaps with estradiol in regulating FSH production. Therefore, the relative role of estradiol vs. the inhibins has also been difficult to determine (discussed further below). The distinct secretory patterns of inhibins A and B may be regulated, at least in part, by the maturational stage of granulosa cell development. In vivo, FSH administration stimulates both inhibin A and B secretion in the early follicular phase from small antral follicles. In vitro, cultured granulosa cells from this stage secrete inhibin A in response to both FSH and cAMP but not inhibin B [35], as well as in response to FSH or LH

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treatment in vivo [36,37]. The inhibin β promoter does not have a cAMP response element which might explain this lack of direct stimulation [38,39]. Several studies suggest that other TGFβ family members, such as activins, BMPs and TGFβ contribute to upregulating inhibin βB subunit production and thus, inhibin B production and secretion [40,41]. In fact, early in the follicular phase when inhibin B rises, an expanding cohort of small antral follicles is growing in response to FSH, suggesting that increased follicle number and their contained expanding and maturing granulosa cells accounts for increased inhibin B levels during the follicular phase in concurrence with FSH. Inhibin B then suppresses FSH via negative feedback at the pituitary, resulting in selection of a single dominant follicle each cycle [18]. Inhibin α and inhibin/activin βA and βB subunit mRNAs and proteins are expressed within granulosa cells at various stages of development. The regulation of α-β dimerization to form inhibin versus β-subunits dimerizing to form activins A, B, or AB remains incompletely understood. One critical factor is the overall level of α-subunit synthesis which is regulated largely by cAMP since increased α-subunit production favors α-β dimers of inhibin [9,42]. In addition, mature α-subunit is glycosylated and elimination of this carbohydrate chain inhibited inhibin secretion without altering activin while mutating β-subunit sequence to incorporate new glycosylation sites also favored formation of inhibin, collectively suggesting that overall glycosylation levels also regulate inhibin versus activin formation [43]. Activin itself can stimulate both α and β subunit synthesis suggesting that further internal feedback amplification once activin is produced [44]. In addition, BMP4 and 7 increase activin A expression and production in human granulosa cells [45]. Thus, regulation of inhibin/activin subunit synthesis is both complex and multifactorial, which accounts for the complicated patterns of these hormones in both animal and human studies. While inhibin is clearly an endocrine hormone with primary actions regulating FSH biosynthesis and release from the pituitary, activin is more likely to be an autocrine/paracrine acting hormone [31]. Within the ovary, activin has been ascribed a number of activities including regulation of germ cell development, primordial follicle assembly and activation (see below), regulation of follicle growth and maturation, promotion of granulosa cell proliferation, suppression of thecal androgen production, and promotion of FSH receptor expression and response [46]. Activin upregulates cyp26b1, which is a retinoic degrading enzyme, a process that regulates sperm mitosis in males [47]. Activin also regulates estrogen receptor expression [48]. In human granulosa-lutein cells from in vitro fertilization (IVF) hyperstimulation, activin promoted expression of P450 aromatase, FSH receptor, and

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estradiol production while inhibiting StAR, LH receptor, and progesterone levels via Smad2 and 3 phosphorylation [49,50]. The autocrine/paracrine roles of activins within the ovary were more fully delineated using inducible CreLox technology to selectively eliminate activin A or activin B in the adult. This was necessary since embryonic deletion of the βA gene resulted in pups that died within 24 h of birth due to numerous developmental defects [51]. Although germ-line βB deficient mice survived to adulthood, offspring died perinatally perhaps due to insufficient lactation [52]. Granulosa cell-specific deletion of the βA gene resulted in subfertility due to enhanced corpus luteum survival that reduced and eventually eliminated estrous cycling [53]. When these mice were crossed with germ-line βB deletion mice, the resulting double knockout females were completely infertile and also accumulated large number of corpus lutea, suggesting that there is substantial redundancy between the βA and βB genes in terms of ovarian function [53]. Using the same strategy, conditional knockout of Smads 2 and 3 in ovarian granulosa cells individually had little detectable effect on ovarian physiology and overall fertility while the combined Smad2/3 knockout dramatically reduced fertility via disrupted follicular development, ovulation, and cumulus cell expansion of ovulated oocytes suggesting that these two second messengers have redundant functions in mediating activin signaling in the ovary [54]. Finally, knocking out the Smad4 common Smad that is required for complexing with phosphorylated Smads to transduce an activin signal produced a similar phenotype of subfertility and multiple defects in folliculogenesis including altered steroidogenesis [55]. These ovaryspecific knockout models provide important clues to the roles of activins within the ovary to regulate follicle formation, development, and maturation. Regulation of follistatin biosynthesis is incompletely understood and may differ depending on the tissue source, but for the most part activin, acting via Smad2 or 3 and FoxL2 appears to be a major simulator of follistatin biosynthesis in pituitary [56–58]. Pathways regulating follistatin synthesis in the ovary include protein kinases A and C [59], BMP2, and FoxL2 [60] as well as wnt/b-catenin [57,61] and Nrf2 [62]. Importantly, follistatin is synthesized as two mRNA transcripts, which produce three distinct follistatin proteins comprising 288, 303, and 315 amino acids in the mature protein [34]. Moreover, the FST288 protein binds tightly to heparin sulfate and thus, remains closely associated with cell surfaces within tissues while the FST315 form does not and is found mainly in the circulation [27,63]. The FST303 protein is mostly found in ovarian follicular fluid [34] and appears to be the dominant form acting in the ovary [18]. A recent study identified the liver as the source for circulating follistatin in humans [64].

The physiological functions of follistatin in the ovary are difficult to dissect given this complexity in biochemistry and the wide distribution of its synthesis within the body. Moreover, germ-line knockout of the follistatin gene resulted in neonatal lethality precluding identification of possible roles. Using the conditional knockout approach, however, follistatin was deleted from granulosa cells in adult ovaries which produced fertility defects including reduced litter number and size leading ultimately to infertility [65]. There was an overall reduction in follicle number along with elevated levels of serum FSH and LH. This study suggested that follistatin plays a key role in regulating follicle development and number and that when compromised, follicles eventually fail to develop and ovulate. Moreover, follistatin dysfunction might contribute to human infertility conditions, such as premature ovarian failure/primary ovarian insufficiency given the similarity of this condition to the phenotype of the granulosa cell-specific follistatin KO. Studies in which the follistatin gene has been inactivated [65,66] do not address the relative roles of the different follistatin isoforms. To determine whether follistatin isoforms have distinct biological actions their properties were assessed individually [27]. Follistatin 288 (FST288) was found to be superior at regulating activin when expressed endogenously and bound to the cell surface compared to FST315 that did not bind to the cell surface [27]. This was true for activin whether applied exogenously or expressed from a transgene endogenously [27]. These observations suggested that FST288 may be sufficient for normal embryonic development given that animals with deletion of all follistatin isoforms were nonviable [66]. In addition, since the FST303 isoform is derived via C-terminal proteolytic processing of the full-length FST315 isoform, elimination of the alternative FST315 mRNA would lead to simultaneous loss of the FST303 ovarian isoform [34] and the circulating FST315 isoform [63] since it is derived by proteolytic processing of the FST315 tail [34]. To test this hypothesis, FST288-only mice were created by removing the alternative splicing sequences that create FST315 transcripts [67]. These mice were born in normal Mendelian ratios verifying that FST288 alone was indeed sufficient for development to adulthood in mice. Interestingly, these mice developed a fertility defect characterized by reduced litter size and frequency, reduced numbers of follicles, and early termination of fertility [67], similar to the granulosa-specific follistatin knockout [65] and to human premature ovarian failure/primary ovarian insufficiency. Detailed follicle counting demonstrated a larger number of primordial, primary and secondary follicles at 8.5 days and a reduced number of healthy antral follicles at 100 and 250 days in FST-288only females demonstrating a more rapid demise of remaining primordial/primary follicles that accounted

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for the early cessation of fertility [67]. In a follow-up study, these females demonstrated that the process of germ cell nest breakdown was extended in duration while apoptosis of forming antral follicles was reduced so that these females were born with more germ cells within the nests [68]. Taken together these studies demonstrate that follistatin is critical for regulating the process of follicle formation and activation, alteration of which leads to reduced fertility and early loss of reproduction. The critical importance of activin signaling in regulating this process of germ cell formation, nest breakdown, follicle formation, and follicle activation/maturation has been amply demonstrated in both mouse and human ovaries [69–71]. Taken together, these studies collectively demonstrate that one critical role of follistatin in the ovary is to regulate activin-mediated germ cell formation, development, and maturation.

HUMAN PHYSIOLOGY AND CLINICAL IMPLICATIONS FSH Regulation for Controlled Follicle Development Changes in inhibin secretion are critical for the precise regulation of FSH and development of a single dominant follicle in each menstrual cycle. FSH levels begin to decline approximately 1 day before peak inhibin B levels in the follicular phase. A time-lag analysis demonstrates an inverse relationship between FSH and inhibin B levels, suggesting that inhibin B may be the most proximate regulator of FSH at this time in the cycle [72]. Evidence for the negative feedback role of inhibin B also comes from the selective FSH rise that occurs in the early follicular phase during normal reproductive aging. The rise in follicular-phase FSH levels is associated with decreased inhibin B levels that result from the decreased small antral follicle number that occurs in ovarian aging [72,73]. The decrease in inhibin B levels occurs earlier than changes in estradiol and inhibin A, the other granulosa cell factors that control FSH, suggesting that the decrease in inhibin B is responsible for the FSH increase during aging [73–76]. These data provide indirect evidence for a role of inhibin B in FSH restraint. In the late luteal phase, the decline in inhibin A levels coincident with the rise in FSH levels across the luteal follicular transition also suggest a role for inhibin A in FSH negative feedback. Inhibin A infusions in the luteal phase suppressed FSH and prevented the FSH rise at menses in the nonhuman primate, providing direct evidence that inhibin A is an important feedback regulator [77]. Similarly, daily inhibin A injections in the follicular phase in the nonhuman primate suppressed bioactive FSH levels [78]. On the contrary, maintenance of a midluteal phase

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estradiol level in women prevented the FSH rise at the luteal-follicular transition despite a decrease in inhibin A levels [79], data have continued to cloud the importance of inhibin A as a negative regulator of FSH in women. Taken together, the evidence suggests inhibin B, and possibly inhibin A, contribute to the negative feedback which regulate FSH levels in the follicular phase of the menstrual cycle.

Ovarian Aging, Perimenopause and Menopause Study of the menopause transition provides support for the role of inhibin B as a marker of ovarian aging and time to menopause in addition to evidence supporting its role as a negative regulator of FSH levels. The stages of reproductive aging have been categorized from early reproductive age through late post menopause based on menstrual cycle regularity, symptoms and hormone levels [80]. These easily identifiable stages allow prediction of time to menopause [80]. The stages are called the STRAW stages based on the Stages of Reproductive Aging Workshop at which they were defined [80]. Several studies demonstrated that inhibin B levels decrease across the STRAW stages of reproductive aging [81]. Therefore, inhibin B levels, along with AMH and antral follicle count, have been included in the STRAW staging system for the prediction of menopause [80]. In addition to its role as a marker of ovarian function across the menopause transition, inhibin may have effects on bone turnover in the perimenopause. Mouse studies and human cell line studies support a direct role for inhibin in suppressing osteoblastogenesis and osteoclastogenesis [82,83]. The effect occurs through blockade of activin and by blocking the stimulatory effects of bone morphogenic proteins (BMPs) [82]. These local effects are consistent with serum inhibin levels, which correlate inversely with markers of bone turnover and formation. For example, decreased inhibin levels are consistent with the increased bone turnover that occurs in the perimenopause [83].

Primary Ovarian Insufficiency Given the known inhibin B decrease in the menopause transition, inhibin B has been evaluated as a marker for primary ovarian insufficiency (POI), previously termed premature ovarian failure. The condition is caused by premature loss of oocytes and follicles, resulting in low estradiol levels, amenorrhea and elevated FSH levels. One of the more common causes of POI results from an expansion in a CGG repeat in the 50 untranslated region of the FMR1 gene, termed a premutation. Inhibin B levels are lower in women who carry the premutation than in control women of the same age even before menstrual

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cycles become irregular, suggesting that they have early ovarian aging [84]. A decrease in inhibin B is clearly seen in women with primary ovarian insufficiency caused by all etiologies, but its ability to identify women with impending POI is not greater than that of AMH [85]. Indeed, AMH appears to be a more sensitive marker of imminent ovarian failure [85]. However, inhibin B levels are particularly important in ovarian insufficiency caused by autoimmunity. Adrenal insufficiency and/or the presence of adrenal antibodies are demonstrated in a majority of women with proven autoimmune oophoritis because the autoimmune reaction targets enzymes common to adrenal and ovarian steroidogenic cells [86]. In early stages of primary ovarian insufficiency caused by autoimmunity, a specific lymphocytic infiltrate invades the steroidogenic theca cells of the follicle, with relative sparing of the granulosa cells [86,87]. In these early stages of autoimmune oophoritis, estradiol production is compromised by the lack of androstenedione and testosterone precursor from theca cells [88]. The elevated FSH levels, in the absence of estradiol negative feedback, continue to drive granulosa cell division and follicle growth with a rise in inhibin B [88]. Thus, high inhibin B levels in the presence of very low estradiol levels serve as an excellent marker of autoimmune oophoritis in the early stages of development [88,89]. Eventually, the lymphocytic infiltrate invades the structures surrounding the theca cells, causing greater follicle destruction and primary ovarian insufficiency.

Puberty In human female neonates, there is a self-limited rise in gonadotropins that stimulates increased inhibin A and B levels for approximately 6 months [90]. Inhibin A levels then decrease to undetectable levels at 6 months after birth, whereas inhibin B levels decrease to detectable but low prepubertal levels. Inhibin B levels are slightly higher from age 6 to 10 years but it is not until approximately age 10 years, in the years leading up to puberty, that inhibin B levels begin a more profound rise [91]. Anchoring inhibin levels to breast development in girls aged 6–11 years demonstrates that inhibin B levels increase gradually from Tanner stage I to II breast development, then increase more sharply to peak at Tanner stage III, indicating the maximum increase in follicle activity across reproductive age [92,93]. The peak inhibin B levels then fall slightly and plateau at stages IV and V as ovulatory menstrual cycles are established [92,93]. Inhibin B levels are correlated with those of FSH and LH levels from Tanner stage 1 through III breast development [93]. In contrast to inhibin B, inhibin A levels increase most dramatically at Tanner stage IV and are inversely correlated with FSH levels, indicating the onset

of ovulatory cycles with corpus luteum activity. Taken together, the prepubertal and pubertal patterns of increase in inhibin B and A mark gonadotropindependent follicle development. Based on the ability to mark follicle activity in puberty, inhibin B has been investigated as a biomarker of abnormal pubertal development. In women with delayed puberty from isolated hypogonadotropic hypogonadism, inhibin B levels were significantly lower than in control subjects in the early follicular phase, consistent with a smaller number of antral follicles. Inhibin B also has excellent sensitivity and specificity to distinguish hypogonadotropic hypogonadism from constitutional delay of growth as a cause of delayed puberty [94]. Inhibin B may serve as a useful indicator of pubertal progression and potential fertility in girls with Prader Willi syndrome, with utility as a clinical marker of the need for contraception [95]. Finally, inhibin B has been evaluated as a marker for precocious puberty although it was unable to differentiate premature thelarche and precocious puberty [96].

Pregnancy and Labor Inhibin A, activin A, and follistatin rise across pregnancy to reach maximum levels at 36 weeks [97]. All are expressed in the placenta. Inhibin A is also produced from the corpus luteum, which is maintained by hCG stimulation from the trophoblast. Activin A rises at the end of pregnancy as a product of the trophoblast and positively correlates with the onset of labor [98]. For unclear reasons, inhibin A is elevated in pregnancies with Down syndrome and has been used for second trimester Down syndrome screening. A recent Cochrane analysis suggests that the small number of studies that used second trimester total hCG, unconjugated estriol (uE3), alpha fetoprotein (AFP), and inhibin A in combination had greater diagnostic accuracy compared to other test combinations that involved only one serum marker or nuchal translucency on ultrasound in the first trimester [99]. Inhibin A and activin A have been evaluated as early second trimester markers of preeclampsia. However, neither has the sensitivity nor specificity that is necessary for use in clinical practice [100]. Inhibin A and activin A have also been evaluated in multimarker panels to identify the location of a pregnancy, whether intrauterine or ectopic, with activin A more promising as a marker of ectopic pregnancy [101]. However, none of the marker panels are yet accurate enough for clinical use.

In Vitro Fertilization and Ovarian Follicle Reserve Evidence that inhibin B derives from the developing cohort of follicles in the menstrual cycle prompted its

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evaluation as a prognostic marker of follicle development during assisted reproductive technologies. Basal prestimulation and early stimulation levels that predict subsequent ovarian responsiveness and pregnancy provide the most useful predictive information. Initial studies using basal inhibin B levels were promising and the inhibin B response to an exogenous FSH ovarian reserve test was predictive of ovarian reserve [102]. However, the majority of studies do not find baseline or stimulated inhibin B levels useful as a screening test for assisted reproductive technologies [103,104]. There is a great deal of overlap in inhibin B levels in subjects who have a good response to treatment compared to those who do not [105]. Taken together, baseline inhibin B and inhibin B during stimulated screening tests do not match the clinical utility of the more easily measured AMH levels. Ovarian markers are also needed as predictors of ovarian function after chemotherapy. Recovery of ovarian function depends on age, the type of chemotherapy, and the pretreatment ovarian reserve. In some studies, inhibin B in combination with AMH predicts ovarian function after chemotherapy for breast cancer [106,107]. Further evaluation of its use as a predictor is necessary.

Ovarian Cancer The majority of ovarian cancers, approximately 90%, derive from ovarian surface epithelium, while 5%–10% arises from granulosa cells. As the inhibins are produced by granulosa cells, the secretion of inhibins is greatest from granulosa cell tumors and inhibin serves as an important marker [108]. Serum inhibin levels are also elevated in mucinous ovarian tumors [108]. A study comparing inhibin levels using a variety of inhibin assays demonstrated that inhibin assays directed to the α subunit of inhibin, which measures both free α subunit and dimeric inhibin, detected 100% of granulosa cell tumors and 70%–90% of mucinous epithelial ovarian carcinomas [108]. The dimeric inhibin B assay also detected over 90% of granulosa cell tumors, but was less useful for detecting mucinous epithelial ovarian tumors [108]. Finally, the utility of the α inhibin subunit directed assay and dimeric inhibin in monitoring granulosa cell tumor recurrence has been established for both juvenile and adult granulosa cell tumors [109,110]. In other ovarian cancers derived from the ovarian surface epithelium, inhibin is less commonly secreted. Total inhibin α subunit directed assays are better than the other inhibin assays at detecting the serous epithelial ovarian tumor subtypes and miscellaneous ovarian tumors although CA125 is superior to inhibin in these cases [108]. A combination of CA125 and an α inhibin subunit directed assay increases the sensitivity of detection slightly but has not been used clinically [108].

In addition to serving as a marker for malignant sexcord stromal tumors, dimeric inhibin secretion can cause unusual menstrual cycle disruptions in cases of benign sex-cord stromal tumors. Dimeric inhibin secretion has been demonstrated from benign sex cord stromal tumors, such as fibrothecomas [111]. These tumors may result in irregular menses or amenorrhea in a premenopausal woman or lower than expected FSH levels in a postmenopausal woman as a consequence of the FSH suppression by dimeric inhibin. Tumor removal can restore menstrual regularity in these rare cases in reproductive age women.

Polycystic Ovary Syndrome PCOS is a disorder defined by two out of three criteria: irregular menses, hyperandrogenism, and polycystic ovary morphology on ultrasound. Follicles are arrested at a small antral stage of development. Although the number of follicles in polycystic ovaries is increased compared to normal ovaries, the majority of studies examining serum inhibin levels in women with PCOS have failed to demonstrate increased inhibin A or inhibin B levels [112,113]. These findings suggest that inhibin B is relatively decreased per follicle in women with PCOS. Consistent with the hypothesis, follicular fluid inhibin A and inhibin B levels are lower in women with PCOS compared to control women [114]. When hormonal and metabolic factors were examined in relation to inhibin levels in women with PCOS, inhibin B levels, but not inhibin A levels, were inversely correlated with BMI and factors related to BMI including LH, insulin, and SHBG [112,113]. As expected based on the correlations, short-term insulin suppression with diazoxide increased inhibin B and hCG administration suppressed inhibin B secretion [112,115]. Recent genome-wide association studies identified a gene variant near the FSHβ gene that is associated with PCOS, lower FSH and higher LH levels [116]. Thus, the slightly lower FSH or higher LH levels may explain the lower inhibin B levels in PCOS. Activin and follistatin levels also demonstrate abnormalities in PCOS. In general, follistatin levels are increased in women with PCOS [117,118]. Activin, in contrast, has been reported as decreased or not different [118]. There is no direct evidence that activin plays a role in the pathophysiology of PCOS. However, the possibility that follistatin plays a role in the pathophysiology is still under investigation.

SUMMARY Now that the genome has been sequenced we know that no additional inhibin/activin/follistatin genes will be discovered. Therefore, future progress will most likely

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come from a better understanding of the physiological roles of each of these proteins both in local, autocrine/ paracrine actions and in endocrine activities. For example, are the different molecular weight forms of dimeric inhibins equal in activity at the pituitary? Do they have different roles within the ovary? Is there any endocrine role for serum activin and follistatin or does the complex in serum merely represent hormones on their way to disposal? Is there a physiological mechanism that can release activin from follistatin since natural dissociation of the complex is essentially nonexistent? How exactly does inhibin block activin signaling? How does the relatively small concentration of inhibin in serum regulate activin action at the pituitary when it binds to the activin receptor with affinities more than 10-fold lower than would be expected for the known inhibin concentration? Are there accessory proteins that have not yet been identified to account for this discrepancy? Clinically, inhibin A serves as a standard marker for Down syndrome in the second trimester of pregnancy. Inhibin α subunit has excellent utility as a biomarker of granulosa cell and mucinous ovarian tumors. Inhibin B levels mark active autoimmune oophoritis at its early stages, before the onset of primary ovarian insufficiency. In contrast, inhibin B levels seemed promising as a fertility marker and as a marker of polycystic ovary morphology. However, AMH levels are superior for assessment of the follicle complement. Prospective trials will determine the utility of inhibin A and activin A for the diagnosis of ectopic pregnancy, inhibin B for the diagnosis of hypogonadotropic hypogonadism and inhibin α subunit as a marker of mucinous ovarian tumor recurrence. Thus, the inhibin field has matured substantially. However, there is still a long way to go to fully understand ovarian physiology and clinical utility of the inhibin family of proteins.

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[65] Jorgez CJ, Klysik M, Jamin SP, Behringer RR, Matzuk MM. Granulosa cell-specific inactivation of follistatin causes female fertility defects. Mol Endocrinol 2004;18:953–67. [66] Matzuk MM, Lu N, Vogel HJ, Sellheyer K, Roop DR, Bradley A. Multiple defects and perinatal death in mice deficient in follistatin. Nature 1995;374:360–3. [67] Kimura F, Sidis Y, Bonomi L, Xia Y, Schneyer A. The follistatin-288 isoform alone is sufficient for survival but not for normal fertility in mice. Endocrinology 2010;151(3):1310–9. [68] Kimura F, Bonomi LM, Schneyer AL. Follistatin regulates germ cell nest breakdown and primordial follicle formation. Endocrinology 2011;152(2):697–706. [69] Bristol-Gould S, Kreeger P, Selkirk C, et al. Postnatal regulation of germ cells by activin: The establishment of the initial follicle pool. Dev Biol 2006;298:132–48. [70] Lengil T, Gancz D, Gilboa L. Activin signaling balances proliferation and differentiation of ovarian niche precursors and enables adjustment of niche numbers. Development 2015;142(5):883–92. [71] Wang Z, Niu W, Wang Y, et al. Follistatin288 regulates germ cell cyst breakdown and primordial follicle assembly in the mouse ovary. PLoS ONE 2015;10(6):e0129643. [72] Robertson DM, Hale GE, Jolley D, Fraser IS, Hughes CL, Burger HG. Interrelationships between ovarian and pituitary hormones in ovulatory menstrual cycles across reproductive age. J Clin Endocrinol Metab 2009;94:138–44. [73] Welt CK, McNicholl DJ, Taylor AE, Hall JE. Female reproductive aging is marked by decreased secretion of dimeric inhibin. J Clin Endocrinol Metab 1999;84:105–11. [74] Klein NA, Houmard BS, Hansen KR, Woodruff TK, Sluss PM, Bremner WJ, Soules MR. Age-related analysis of inhibin A, inhibin B, and activin a relative to the intercycle monotropic folliclestimulating hormone rise in normal ovulatory women. J Clin Endocrinol Metab 2004;89:2977–81. [75] Burger HG, Dudley EC, Robertson DM, Dennerstein L. Hormonal changes in the menopause transition. Recent Prog Horm Res 2002;57:257–75. [76] Reame NE, Wyman TL, Phillips DJ, de Kretser DM, Padmanabhan V. Net increase in stimulatory input resulting from a decrease in inhibin B and an increase in activin A may contribute in part to the rise in follicular phase follicle-stimulating hormone of aging cycling women. J Clin Endocrinol Metab 1998;83:3302–7. [77] Stouffer RL, Dahl KD, Hess DL, Woodruff TK, Mather JP, Molskness TA. Systemic and intraluteal infusion of inhibin A or activin A in rhesus monkeys during the luteal phase of the menstrual cycle. Biol Reprod 1994;50:888–95. [78] Molskness TA, Woodruff TK, Hess DL, Dahl KD, Stouffer RL. Recombinant human inhibin-A administered early in the menstrual cycle alters concurrent pituitary and follicular, plus subsequent luteal, function in rhesus monkeys. J Clin Endocrinol Metab 1996;81:4002–6. [79] Lahlou N, Chabbert-Buffet N, Christin-Maitre S, Le Nestour E, Roger M, Bouchard P. Main inhibitor of follicle stimulating hormone in the luteal-follicular transition: inhibin A, oestradiol, or inhibin B? Hum Reprod 1999;14:1190–3. [80] Harlow SD, Gass M, Hall JE, Lobo R, Maki P, Rebar RW, Sherman S, Sluss PM, de Villiers TJ, Group SC. Executive summary of the stages of reproductive aging workshop + 10: addressing the unfinished agenda of staging reproductive aging. Fertil Steril 2012;97:843–51. [81] Hale GE, Zhao X, Hughes CL, Burger HG, Robertson DM, Fraser IS. Endocrine features of menstrual cycles in middle and late reproductive age and the menopausal transition classified according to the staging of reproductive aging workshop (STRAW) staging system. J Clin Endocrinol Metab 2007;92:3060–7. [82] Gaddy-Kurten D, Coker JK, Abe E, Jilka RL, Manolagas SC. Inhibin suppresses and activin stimulates osteoblastogenesis and

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osteoclastogenesis in murine bone marrow cultures. Endocrinology 2002;143:74–83. Perrien DS, Achenbach SJ, Bledsoe SE, Walser B, Suva LJ, Khosla S, Gaddy D. Bone turnover across the menopause transition: correlations with inhibins and follicle-stimulating hormone. J Clin Endocrinol Metab 2006;91:1848–54. Welt CK, Smith PC, Taylor AE. Evidence of early ovarian aging in fragile X premutation carriers. J Clin Endocrinol Metab 2004;89:4569–74. Knauff EA, Eijkemans MJ, Lambalk CB, ten Kate-Booij MJ, Hoek A, Beerendonk CC, Laven JS, Goverde AJ, Broekmans FJ, Themmen AP, de Jong FH, Fauser BC. Dutch premature ovarian failure C. Anti-mullerian hormone, inhibin B, and antral follicle count in young women with ovarian failure. J Clin Endocrinol Metab 2009;94:786–92. Bakalov VK, Anasti JN, Calis KA, Vanderhoof VH, Premkumar A, Chen S, Furmaniak J, Smith BR, Merino MJ, Nelson LM. Autoimmune oophoritis as a mechanism of follicular dysfunction in women with 46,XX spontaneous premature ovarian failure. Fertil Steril 2005;84:958–65. Hoek A, Schoemaker J, Drexhage HA. Premature ovarian failure and ovarian autoimmunity. Endocr Rev 1997;18:107–34. Welt CK, Falorni A, Taylor AE, Martin KA, Hall JE. Selective theca cell dysfunction in autoimmune oophoritis results in multifollicular development, decreased estradiol, and elevated inhibin B levels. J Clin Endocrinol Metab 2005;90:3069–76. Tsigkou A, Marzotti S, Borges L, Brozzetti A, Reis F, Candeloro P, Luisa Bacosi M, Bini V, Petraglia F, Falorni A. High serum inhibin concentration discriminates autoimmune oophoritis from other forms of primary ovarian insufficiency. J Clin Endocrinol Metab 2008;93:1263–9. Bergada I, Rojas G, Ropelato G, Ayuso S, Bergada C, Campo S. Sexual dimorphism in circulating monomeric and dimeric inhibins in normal boys and girls from birth to puberty. Clin Endocrinol (Oxf ) 1999;51:455–60. Crofton PM, Evans AE, Groome NP, Taylor MR, Holland CV, Kelnar CJ. Dimeric inhibins in girls from birth to adulthood: relationship with age, pubertal stage, FSH and oestradiol. Clin Endocrinol (Oxf ) 2002;56:223–30. Sims EK, Addo OY, Gollenberg AL, Himes JH, Hediger ML, Lee PA. Inhibin B and luteinizing hormone levels in girls aged 6-11 years from NHANES III, 1988-1994. Clin Endocrinol (Oxf ) 2012;77:555–63. Sehested A, Andersson AM, Muller J, Skakkebaek NE. Serum inhibin A and inhibin B in central precocious puberty before and during treatment with GnRH agonists. Horm Res 2000;54:84–91. Binder G, Schweizer R, Haber P, Blumenstock G, Braun R. Accuracy of endocrine tests for detecting hypogonadotropic hypogonadism in girls. J Pediatr 2015;167:674–8 [e671]. Eldar-Geva T, Hirsch HJ, Pollak Y, Benarroch F, Gross-Tsur V. Management of hypogonadism in adolescent girls and adult women with Prader-Willi syndrome. Am J Med Genet A 2013;161A:3030–4. De Filippo G, Rendina D, Nazzaro A, Lonardo F, Bouvattier C, Strazzullo P. Baseline inhibin B levels for diagnosis of central precocious puberty in girls. Horm Res Paediatr 2013;80:207–12. Fowler PA, Evans LW, Groome NP, Templeton A, Knight PG. A longitudinal study of maternal serum inhibin-A, inhibin-B, activin-A, activin-AB, pro-alphaC and follistatin during pregnancy. Hum Reprod 1998;13:3530–6. Woodruff TK, Sluss P, Wang E, Janssen I, Mersol-Barg MS, Activin A. Follistatin are dynamically regulated during human pregnancy. J Endocrinol 1997;152:167–74. Alldred SK, Takwoingi Y, Guo B, Pennant M, Deeks JJ, Neilson JP, Alfirevic Z. First and second trimester serum tests with and without first trimester ultrasound tests for Down’s syndrome screening. Cochrane Database Syst Rev 2017;3:CD012599.

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[100] Wu P, van den Berg C, Alfirevic Z, O’Brien S, Rothlisberger M, Baker PN, Kenny LC, Kublickiene K, Duvekot JJ. Early pregnancy biomarkers in pre-eclampsia: a systematic review and metaanalysis. Int J Mol Sci 2015;16:23035–56. [101] Senapati S, Sammel MD, Butts SF, Takacs P, Chung K, Barnhart KT. Predicting first trimester pregnancy outcome: derivation of a multiple marker test. Fertil Steril 2016;106:1725–32: e1723. [102] Kwee J, Schats R, McDonnell J, Themmen A, de Jong F, Lambalk C. Evaluation of anti-Mullerian hormone as a test for the prediction of ovarian reserve. Fertil Steril 2008;90:737–43. [103] Hall JE, Welt CK, Cramer DW. Inhibin A and inhibin B reflect ovarian function in assisted reproduction but are less useful at predicting outcome. Hum Reprod 1999;14:409–15. [104] Jayaprakasan K, Campbell B, Hopkisson J, Johnson I, RaineFenning N. A prospective, comparative analysis of anti-Mullerian hormone, inhibin-B, and three-dimensional ultrasound determinants of ovarian reserve in the prediction of poor response to controlled ovarian stimulation. Fertil Steril 2010;93:855–64. [105] Muttukrishna S, Suharjono H, McGarrigle H, Sathanandan M. Inhibin B and anti-Mullerian hormone: markers of ovarian response in IVF/ICSI patients? BJOG 2004;111:1248–53. [106] Su HI, Sammel MD, Green J, Velders L, Stankiewicz C, Matro J, Freeman EW, Gracia CR, DeMichele A. Antimullerian hormone and inhibin B are hormone measures of ovarian function in late reproductive-aged breast cancer survivors. Cancer 2010;116:592–9. [107] Anders C, Marcom PK, Peterson B, Gu L, Unruhe S, Welch R, Lyons P, Behera M, Copland S, Kimmick G, Shaw H, Snyder S, Antenos M, Woodruff T, Blackwell K. A pilot study of predictive markers of chemotherapy-related amenorrhea among premenopausal women with early stage breast cancer. Cancer Invest 2008;26:286–95. [108] Robertson DM, Stephenson T, Pruysers E, McCloud P, Tsigos A, Groome N, Mamers P, Burger HG. Characterization of inhibin forms and their measurement by an inhibin alpha-subunit ELISA in serum from postmenopausal women with ovarian cancer. J Clin Endocrinol Metab 2002;87:816–24. [109] Geerts I, Vergote I, Neven P, Billen J. The role of inhibins B and antimullerian hormone for diagnosis and follow-up of granulosa cell tumors. Int J Gynecol Cancer 2009;19:847–55. [110] Wu H, Pangas SA, Eldin KW, Patel KR, Hicks J, Dietrich JE, Venkatramani R. Juvenile granulosa cell tumor of the ovary:

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a clinicopathologic study. J Pediatr Adolesc Gynecol 2017;30: 138–43. van Liempt SW, van Rheenen-Flach LE, van Waesberghe JH, Bleeker MC, Piek JM, Lambalk CB. Solely inhibin B producing ovarian tumour as a cause of secondary amenorrhoea with hot flushes: case report and review of literature. Hum Reprod 2012;27:1144–8. Welt CK, Taylor AE, Martin KA, Hall JE. Serum inhibin B in polycystic ovary syndrome: regulation by insulin and luteinizing hormone. J Clin Endocrinol Metab 2002;87:5559–65. Cortet-Rudelli C, Pigny P, Decanter C, Leroy M, MaunouryLefebvre C, Thomas-Desrousseaux P, Dewailly D. Obesity and serum luteinizing hormone level have an independent and opposite effect on the serum inhibin B level in patients with polycystic ovary syndrome. Fertil Steril 2002;77:281–7. Welt CK, Taylor AE, Fox J, Messerlian GM, Adams JM, Schneyer AL. Follicular arrest in polycystic ovary syndrome is associated with deficient inhibin A and B biosynthesis. J Clin Endocrinol Metab 2005;90:5582–7. Shayya RF, Rosencrantz MA, Chuan SS, Cook-Andersen H, Roudebush WE, Irene Su H, Shimasaki S, Chang RJ. Decreased inhibin B responses following recombinant human chorionic gonadotropin administration in normal women and women with polycystic ovary syndrome. Fertil Steril 2014;101:275–9. Day FR, Hinds DA, Tung JY, Stolk L, Styrkarsdottir U, Saxena R, Bjonnes A, Broer L, Dunger DB, Halldorsson BV, Lawlor DA, Laval G, Mathieson I, McCardle WL, Louwers Y, Meun C, Ring S, Scott RA, Sulem P, Uitterlinden AG, Wareham NJ, Thorsteinsdottir U, Welt C, Stefansson K, Laven JS, Ong KK, Perry JR. Causal mechanisms and balancing selection inferred from genetic associations with polycystic ovary syndrome. Nat Commun 2015;6:8464. Norman RJ, Milner CR, Groome NP, Robertson DM. Circulating follistatin concentrations are higher and activin concentrations are lower in polycystic ovarian syndrome. Hum Reprod 2001;16:668–72. Teede H, Ng S, Hedger M, Moran L. Follistatin and activins in polycystic ovary syndrome: relationship to metabolic and hormonal markers. Metabolism: clinical and experimental 2013;62:1394–400.

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C H A P T E R

7 Cell-Cell Interactions in Ovarian Follicles: Role of TGF-β Superfamily Members Hsun-Ming Chang, Yi-Min Zhu, Peter C.K. Leung INTRODUCTION Human ovarian follicles are the functional units of the female reproductive system that develop through different stages, including primordial, primary, secondary, antral, and peri-ovulatory follicle stages, and subsequently ovulate in response to the LH surge. Experiments using in vitro culture techniques have demonstrated that the oocyte and follicular cells, including granulosa cells (GCs) and theca cells (TCs), interact with each other to achieve full ovulatory capacity, which is strictly dependent on bidirectional communication [1]. This two-way communication between the germ cells and somatic cells is accomplished by either cell-cell contact or paracrine factors produced by the neighboring cells [2]. Dysregulation of follicular development may impair female reproductive function and lead to several gynecological and endocrine diseases, such as polycystic ovary syndrome (PCOS), primary ovarian insufficiency, chronic anovulation, and even ovarian cancers [3–5]. Thus, follicular transition and development are regulated by a highly coordinated process that involves multiple control systems, including neuronal, neuroendocrinal, endocrinal, paracrinal, and autocrinal systems [6]. In these control systems, the principal roles of GnRH, the pituitary gland-derived gonadotropins, FSH and LH, and the gonadal hormones (androgen, estrogen, and progesterone) in regulating reproductive functions have been well established. However, normal reproductive and follicular functions also rely on various locally produced autocrine and paracrine signals, cytokines, and growth factors [7]. For several years, great effort has been devoted to the study of these intraovarian regulators and how they affect folliculogenesis. In vitro and animal studies have demonstrated that transforming growth factor-β (TGF-β) superfamily members, which include

The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00007-8

activins/inhibins, TGF-βs, bone morphogenetic proteins (BMPs), growth differentiation factors (GDFs), and anti-M€ ullerian hormone (AMH), play critical roles in the formation and regulation of germ cells, follicular development, steroidogenesis, oocyte maturation, and implantation [7,8]. Moreover, natural and experimental gene mutations of TGF-β superfamily ligands and dysregulated TGF-β signaling are associated with several pathological disorders in human reproduction [9]. Much research on the regulation of ovarian function has been performed in a wide range of species. Even though the general aspects of ovarian follicle development are highly similar across species, there is significant species variation with regard to germ cell formation as well as ovarian cell-specific gene expression and regulation. Indeed, most of the relevant data have been generated from studies in rodents. An advanced understanding of the physiology and pathology of human ovaries has only been obtained recently because of emerging technologies including tissue microarrays, recombinant human proteins, pharmaceutical development, new experimental settings, and immortalized human cell lines. Therefore, this chapter predominantly focuses on human oocyte-somatic cell interaction and human ovarian disorders associated with TGF-β superfamily signaling.

TGF-β SUPERFAMILY LIGANDS TGF-β superfamily members are a group of structurally conserved but functionally diverse growth factors that regulate fundamental cellular properties and vital cellular processes, such as proliferation, differentiation, communication, apoptosis, and tissue remodeling [10]. With >40 structurally related proteins, the TGF-β

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7. CELL-CELL INTERACTIONS IN OVARIAN FOLLICLES: ROLE OF TGF-β SUPERFAMILY MEMBERS

superfamily is composed of several subfamilies, including TGF-βs (comprising TGF-β1, TGF-β2, and TGF-β3), BMPs and GDFs (with >20 members), activins (including activin A, B, and AB) and inhibins (inhibin α and β), AMH, the glial cell-derived neurotrophic factors (GDNFs) (including GDNF, artemin, and neurturin), and NODAL [11,12]. These ligands are initially translated as similar structures containing three components: a signal peptide (20–30 amino acids long), a long prodomain, and a mature domain (mature peptide). The inactive TGF-β precursors are further proteolytically cleaved by the proprotein convertase subtilisin/kexin (a family that is composed of nine members, PCSK1-PCSK9) [13]. Several members of this family, such as PCSK3 (also known as furin), can recognize the specific amino acid motif Arg-X-Arg/Lys-Arg- located at the linker site (between the prodomain and mature domain) and induce the formation of mature homodimers or heterodimers [14], which are covalently linked by an interchain disulfide bond between conserved cysteine residues [15]. While the homodimers are the main bioactive proteins that perform cellular actions, several heterodimers of this superfamily, such as TGF-β1/TGF-β2, TGF-β2/TGF-β3, BMP2/BMP7, BMP2/BMP6, BMP4/BMP7, BMP15/ GDF9, inhibin βA/inhibin βB (activin AB), and inhibin βB/inhibin βC (activin AC), have been shown to exist in both in vivo and in vitro systems [16,17]. In general, most of these heterodimers are much more biologically potent than their counterpart homodimers with regard to inducing cellular functions [17,18].

TGF-β SIGNALING RECEPTORS With the exception of the inhibin and GDNF subfamilies, TGF-β superfamily ligands exert their cellular effects by binding to distinct sets of dual transmembrane Ser/ Thr kinase receptors, type I and type II receptors [19]. Dimeric ligands initially bind to the cognate type II receptors, and the complex then recruits and activates type I receptors, ultimately leading to the phosphorylation and activation of R-SMADs (receptor-regulated SMADs) [20]. In mammals, seven distinct type I receptors (also known as activin receptor like-kinase 1–7) (ALK 1–7) and five type II receptors (TβR2, ACVR2A, ACVR2B, BMPR2, and AMHR2) have been identified to induce the assembly of a heterotetrameric complex [11]. In this regard, the sequence and structural differences among the TGF-β superfamily ligands may be responsible for their different binding affinities for the various type II and type I receptors. Moreover, the binding specificities of ligands for type I receptors can be affected by the type II receptor involved [21].

TGF-β SIGNAL TRANSDUCTION PATHWAY Upon ligand-receptor binding, the activation of type I receptors through the phosphorylation of their intracellular kinase domains leads to the phosphorylation of the downstream R-SMAD signal transducers, the transcription factors SMAD1–8 [19]. There are two general R-SMAD activation models: SMAD1/5/8 respond to BMPs, AMH, and some GDFs via ALK2, ALK3, and ALK6, whereas SMAD2/3 respond to TGF-βs, activins, some GDFs and NODAL via ALK4, ALK5, and ALK7 [22]. Finally, activated R-SMADs associate with a common SMAD (Co-SMAD), SMAD4, and this complex further translocates to the nucleus to modulate target gene expression by interacting with other transcription factors in various cell types and tissues [11,20]. However, recent studies have challenged the conventional understanding that BMPs induce cellular action solely via canonical SMAD1/5/8 signaling. A novel noncanonical SMAD2/3 signaling pathway that is commonly activated by TGF-βs and activins were reported to be induced by BMPs to modulate hormone production and cancer progression [23]. Interestingly, our recent studies also showed that BMP-induced upregulation of hyaluronan synthase type 2 expression occurs through the ALK4/5/7-mediated noncanonical SMAD2/3 signaling pathway in human GCs [24]. Aside from the SMAD-dependent signaling pathway, several SMAD-independent (non-SMAD) signaling pathways have been identified to mediate TGF-β superfamily ligands in specific tissues [25]. Among them, mitogen-activated protein kinase (MAPK) signaling is the major SMAD-independent pathway activated by TGF-β ligands. Studies using dominant-negative SMADs or in SMAD4-deficient cells have indicated that MAPK pathway activation occurs in a SMAD signalingindependent manner [26]. It is increasingly apparent that BMPs and TGF-βs can directly activate extracellular signal-regulated kinase (ERK), JNK, p38MAPK, phosphatidylinositol-3-kinase (PI3K), protein kinase (PK) A, PKC, and PKD signaling pathways in various cells [26]. Furthermore, TGF-β-induced activation of ERK or JNK signaling can lead to SMAD phosphorylation and subsequently regulate the activation of the SMAD pathway [27]. In the human ovary, TGF-β1 and GDF8 decrease the expression of steroidogenic acute regulatory protein (StAR) by activating both the SMAD3 and ERK pathways [28,29]. The subsequent signaling pathway activated by ligand-induced receptor recruitment primarily depends on the extracellular environment, neighboring cellular activity, and cross talk with other signaling pathways [30].

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EXPRESSION OF TGF-β SUPERFAMILY LIGANDS IN THE HUMAN OVARY

EXPRESSION OF TGF-β SUPERFAMILY LIGANDS IN THE HUMAN OVARY Although many studies on the expression of TGF-β superfamily ligands have been performed in mammalian ovaries and have provided important information, the most comprehensive studies have focused on adult rats [7]. However, the ovarian expression pattern of TGF-β ligands in rats does not necessarily recapitulate that in humans. For example, both BMP4 and BMP7 are mainly expressed in theca and stroma cells of rat follicles [31], yet they can be detected in human oocytes [32]. Exclusively expressed in rodent and cattle oocytes [33], GDF9 was identified in human GCs and oocytes [34]. Similarly, BMP15, which was previously thought to be oocytederived growth factor, has been detected in human GCs, cumulus cells, and oocytes [35]. Currently, even though extensive research has focused on the functional roles of TGF-β superfamily ligands in the regulation of normal ovarian biology, relatively little is known about TABLE 1

the spatiotemporal expression patterns of these growth factors in the human ovary. Originally identified in the musculoskeletal system, GDF8 has been found to be expressed in the GCs of growing follicles in chicken, bovine, and human ovaries [36,37]. Recent studies have indicated that this unique TGF-β superfamily member is an intraovarian growth factor that plays a critical role in the regulation of human ovarian functions [37]. The cellular localization and follicular fluid concentration of the TGF-β superfamily ligands in human ovarian tissue are listed in Table 1. Positive TGF-β1 immunostaining can be observed in human oocytes and follicle cells (GCs and TCs), and the intensity of immunostaining in the follicle cells increases along with the follicle size [38]. Moreover, both the small and large luteal cells express TGF-β1 in the corpus luteum. However, TGF-β2 is exclusively expressed in human TCs and the small luteal cells of the corpus luteum [38]. Activins and inhibins are disulfide-linked homodimers or heterodimers that contain inhibin α, inhibin βA,

Localization of TGF-β Ligands in the Human Ovary

Ligands

Localization

Expression

Detection method

BMP2

Luteinized GC

mRNA

RT-qPCR

CL

mRNA

RT-qPCR

FF (1–115 ng/mL)

Protein

ELISA

Luteinized GC

mRNA and protein

RT-qPCR, IHC

Protein

IHC

Protein

IHC

GC

Protein

IHC

CL

mRNA and protein

RT-qPCR, IHC

FF (1–20 pg/mL)

Protein

ELISA

BMP5

Luteinized GC

mRNA

RT-qPCR

BMP6

Luteinized GC

mRNA and protein

RT-qPCR, IHC

CL

mRNA and protein

RT-qPCR, IHC

Luteinized GC

mRNA and protein

RT-qPCR, IHC

Oocyte

Protein

IHC

Theca/stroma cells

Protein

IHC

GC

Protein

IHC

FF (50–130 ng/mL)

Protein

ELISA

BMP8A

Luteinized GC

mRNA

RT-qPCR

BMP15

Preantral follicles (>75 μm)

mRNA

Tissue microarray

Cumulus cells

mRNA

RT-qPCR

Oocyte

mRNA and protein

In situ hybridization, RT-qPCR, IHC

BMP4

a

Oocyte

a

Theca/stroma cells a

BMP7

Continued

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110 TABLE 1 Ligands

7. CELL-CELL INTERACTIONS IN OVARIAN FOLLICLES: ROLE OF TGF-β SUPERFAMILY MEMBERS

Localization of TGF-β Ligands in the Human Ovary—cont’d Localization

Expression

Detection method

Theca/stroma cells

mRNA and protein

In situ hybridization, IHC

GC

mRNA and protein

In situ hybridization, IHC

FF

Protein

Western blot

Luteinized GC

mRNA

RT-qPCR

FF (2.01–4.17 ng/mL)

Protein

ELISA

Preantral follicles

mRNA

Tissue microarray

Cumulus cells

mRNA

RT-qPCR

Oocyte

mRNA and protein

In situ hybridization, IHC

Luteinized GC

mRNA and protein

RT-PCR, Western blot

FF (0.83–53.9 ng/mL)

Protein

ELISA

GC of primary, secondary preantral and small antral follicles

Protein

IHC

FF from small antral follicle (57–1124 ng/mL)

Protein

ELISA

FF from preovulatory follicle (50.1–100 ng/mL)

Protein

ELISA

GC, theca/stroma cells, oocyte of small and large follicles

Protein

IHC

Luteal cells (small and large) of CL

Protein

IHC

FF (0.236–18.03 ng/mL)

Protein

RI

Theca/stroma cells

Protein

IHC

Small luteal cells of CL

Protein

IHC

GC and TCs of preovulatory follicles

DNA

Northern blot

GC and TC of small antral follicles

Protein

IHC

Inhibin βA

GC and TC of early, small antral follicles

Protein

IHC

Inhibin βB

GC and TC of small antral follicles

Protein

IHC

Activin A

FF (1.7–267.9 ng/mL)

Protein

ELISA

Inhibin A

FF (7.9–436 ng/mL)

Protein

ELISA

Inhibin B

FF (9.7–786 ng/mL)

Protein

ELISA

FF from small antral follicle (57–200 ng/mL)

Protein

ELISA

GDF8

GDF9

AMH

TGF-β1

TGF-β2

Inhibin α

CL, corpora lutea; ELISA, enzyme-linked immunosorbent assay; FF, follicular fluid; GC, granulosa cell; IHC, immunohistochemistry; RI, radioimmunoassay; RT-qPCR, quantitative real-time polymerase chain reaction; TC, theca cell. a Expression only in human fetal ovaries.

or inhibin βB subunits, and the expression of these inhibin subunits has been detected in normal and PCOS ovaries [39–41]. During follicle development, inhibin α protein is expressed in the GCs and TCs of human small antral follicles, but not in preantral follicles [42]. In addition, inhibin βA and inhibin βB proteins are mainly confined to the GCs and TCs of the adult human ovary [42]. Notably, the co-localization of inhibin α and inhibin βA mRNA in TCs supports the hypothesis that inhibin has an autocrine function in these cells [40]. In adult mammals, AMH is dynamically expressed in the developing ovarian follicles [43]. Expression of AMH in GCs starts shortly after the primordial follicle begins to

grow [44]. In humans, AMH mRNA and protein are detectable in the GCs of preantral follicles, with peak levels (1124  158 ng/mL) found in the GCs of antral follicles approximately 3 mm in diameter, followed by a reduction in the GCs of the preovulatory follicles [45]. However, AMH has also been detected in endometrial and endometriotic tissues [46]. The expression pattern of AMH in the human ovary suggests that this GC-derived growth factor plays a critical role in the regulation of intrafollicular functions during the midfollicular phase at the time of follicular selection. In most species, follicular size is associated with the development of the oocyte and follicle cells, which is

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EXPRESSION OF TGF-β RECEPTORS IN THE HUMAN OVARY

required to trigger a stage-dependent cascade of molecular maturation events in the nucleus and cytoplasm [47]. The maturation of the oocyte during the antral follicular stage is highly dependent on various intraovarian factors present in the follicular fluid [48]. A crucial balance exists between the physiological concentration and duration of these intrafollicular factors that is required to facilitate the development and maturation of oocyte [49]. Therefore, dysregulation of any of these factors may result in abnormal follicular development and ovulation dysfunction in humans [3,49]. Previous clinical studies have indicated that many TGF-β superfamily members are detectable in the follicular fluid, and some of them are associated with oocyte quality (Table 1). For example, the intrafollicular concentrations of BMP2 and BMP15 are correlated with the rates of oocyte fertilization and oocyte maturation [50,51]. The follicular fluid levels of inhibin A and inhibin B are significantly higher in follicles with a recoverable oocyte; however, the levels of both inhibins are not correlated with subsequent oocyte fertilization [52]. The mature form of GDF8 protein is detectable (2.01–4.17 ng/mL) in the follicular fluid, and its concentration is negatively correlated with the concentration of intrafollicular progesterone [53]. The concentration of AMH in the fluid of individual follicles is positively correlated with the serum AMH concentration, and the AMH concentrations in individual follicles do not significantly differ within the same patient [54]. Taken together, the TGF-β superfamily ligands are highly active throughout all follicular stages, including the corpus luteum. These intraovarian growth factors interact and cooperate to play central roles in regulating follicular development, oocyte maturation, and luteal function.

EXPRESSION OF TGF-β RECEPTORS IN THE HUMAN OVARY Functional receptors for TGF-β superfamily ligands have been identified in various follicular compartments of developing follicles in the human ovary [55] (Table 2). In a comparison of the transcripts of five sizematched human preantral follicles, BMPR2 was found to be the most highly expressed type II receptor in these follicles, followed by AMHR2 [55]. During follicular development, BMP2 is highly expressed in primordial and early primary follicles, with a progressive increase in expression throughout the preantral follicular stage, whereas AMHR2 is consistently expressed throughout each follicular stage [55]. By comparison, TβR2, ACVR2A, and ACVR2B are expressed at relatively low levels in human preantral follicles [55]. Among the seven type I receptors, ALK3, ALK4, ALK5, and ALK6 are expressed at a moderate level,

TABLE 2 Localization of TGF-β Type I and Type II Receptors in the Human Ovary Receptors

Localization

Expression

Detection method

ALK2

Luteinized GC

mRNA

RT-qPCR

ALK3

Luteinized GC

mRNA and protein

RT-qPCR, IHC

GC

Protein

IHC

Oocyte

Protein

IHC

CL

mRNA and protein

RT-qPCR, IHC

ALK4

Pre-GC and stroma of fetal ovary

Protein

IHC

ALK5

Luteinized GC

mRNA

RT-qPCR

GC, TC, and interstitial cells of premenopausal follicles

Protein

IHC

Luteinized GC

mRNA and protein

RT-qPCR, IHC

Oocyte

Protein

IHC

Stroma cells

Protein

IHC

GC

Protein

IHC

CL

mRNA and protein

RT-qPCR, IHC

TβR2

GC, TC and interstitial cells of premenopausal follicles

Protein

IHC

ACVR2A

GC and TC of small antral follicles

Protein

IHC

Oocytes of secondary follicles

Protein

IHC

ACVR2B

GC and TC of small antral follicles

Protein

IHC

BMPR2

Luteinized GC

mRNA and protein

RT-qPCR, IHC

Oocyte

Protein

IHC

Stroma cells

Protein

IHC

GC

Protein

IHC

CL

mRNA and protein

RT-qPCR, IHC

AMHR2

GCs of preantral follicles

mRNA

Microarray

TGFβR3

GCs of preantral follicles

mRNA

Microarray

ALK6

CL, corpora lutea; ELISA, enzyme-linked immunosorbent assay; FF, follicular fluid; GC, granulosa cell; IHC, immunohistochemistry; RI, radioimmunoassay; RT-qPCR, quantitative real-time polymerase chain reaction; TC, theca cell.

I. THE OVARIAN FOLLICULAR APPARATUS: OPERATIONAL CHARACTERISTICS

112

7. CELL-CELL INTERACTIONS IN OVARIAN FOLLICLES: ROLE OF TGF-β SUPERFAMILY MEMBERS

and these receptors are all relatively consistently expressed during follicular growth [55]. Moreover, ALK3 and ALK6 have been identified in oocytes from the primordial to antral follicular stages [32]. Compared with these type I receptors, ALK1, ALK2, and ALK7 are generally expressed at low levels in ovarian follicles [55]. Transforming growth factor β type III receptor (TGFβR3) is also known as betaglycan. This specific receptor has been shown to be an accessory co-receptor of TGF-βs. In many cell types, TGFβR3 inhibits TGF-β signaling by preventing the formation of the TGFβR1-TGFβR2 heterocomplex, making it a potent negative regulator of TGF-β [56]. TGFβR3 has also been shown to bind inhibin, and the presence of TGFβR3 enhances the affinity of inhibin for the activin receptor, ACVR2 in the ovary [57]. The high expression of TGFβR3 in human preantral follicles suggests its role in regulating the activity of activin/inhibin during the preantral follicular stage [55].

PRIMORDIAL GERM CELL DEVELOPMENT At the time of gastrulation, a group of germline stem cells that later give rise to gametes, sperms, and oocytes are described as primordial germ cells (PGCs) [58]. These cells initially migrate to the primitive gonadal fold where they mitotically proliferate and increase in number [59]. Subsequently, these cells differentiate into primordial follicles, which include oocytes and the surrounding somatic cells [59]. Members of the TGF-β superfamily differentially regulate gonadal development in humans and rodents based on their spatiotemporal organization [60]. Mouse PGCs are induced from pluripotent epiblast cells by the synergistic action of extraembryonic cell-derived BMP4 and BMP8B [61]. Moreover, visceral endoderm-derived BMP2 increases the numbers of mouse PGCs [61]. During human fetal ovary development, upregulation of MSX2 in response to BMP4 induces cell apoptosis and modulates postmigratory PGC numbers [60]. Before follicle formation, GDF9 is transiently secreted by oocytes in combination with somatic cell-derived activin βA signaling (secreted by somatic cells) to determine the survival of selected germ cells [62]. In human fetal ovaries, the inhibin βA and βB subunits and activin receptors are expressed in oocytes and somatic cells [63]. An in vitro study showed that activin A increased both the number of oogonia and oogonial proliferation, indicating that activin may be involved in the promotion of germ cell proliferation during primordial follicle formation in the human ovary [63].

REGULATION OF OVARIAN PREANTRAL FOLLICLE DEVELOPMENT In the last couple of decades, interest in studying fertility preservation has grown because of the clinical demand for increased numbers of mature oocytes from dormant primordial follicles. Indeed, the underlying mechanisms by which the ovarian follicles develop from the primordial stage to the antral stage are more complex than predicted. During this development, GCs transform from fibroblast-like cells into the cuboidal shape of epithelial cells with intercellular gap junctions, accompanied by progressive remodeling of the basement membrane [64]. These GCs further divide and form layers that are located at different locations (cumulus, mural, or periantral area), with differential future differentiation potential and functions [64]. In this intrafollicular microenvironment, the oocyte, GCs, TCs, and basement membrane are the key components of early follicular development. Preantral follicles are broadly defined as any time or stage prior to the formation of antral fluid in the central cavity (antrum formation). Most information on the regulation of mammalian preantral follicle development has been derived from rodent models. BMPs play a critical role in postnatal ovarian development. Both BMP4 and BMP7 are expressed by the TC layer of developing rat follicles, and BMPR2 receptors are expressed in the oocytes of primary follicles and GCs of developing rat follicles [8]. Studies using rodent ovary culture showed that BMP4 and BMP7 promote the advancement of primary follicles to the secondary follicle stage [65]. Studies using in vivo and organ culture techniques also indicate that GDF9 may induce primordial follicle progression and promote the transition of primary follicles to secondary follicles [66]. Similar to GDF9, BMP15 is restricted to the oocytes of primary and developing rat follicles [67]. BMP15 is not essential for early follicular development, whereas this growth factor has a stimulatory effect on the proliferation of rat GCs [68]. In immature rat ovaries, treatment with the combination of FSH and TGF-β1 reduced the numbers of preantral follicles, indicating an inhibitory role for TGF-β1 (in combination with FSH) in rat follicular development and progression [69]. In the last decade, the importance of AMH in female reproduction has attracted much attention from many research teams. Initially known as a male hormone due to its role in male differentiation, this hormone is produced by the testis and is able to induce the regression of the M€ ullerian duct during fetal sexual differentiation [70]. Subsequent studies showed that AMH is also produced in the GCs of the ovary, and it is a critical inhibitor of primordial follicle recruitment during follicular development [71]. Targeted ablation of the AMH gene in mice promotes the recruitment of primordial follicles, resulting

I. THE OVARIAN FOLLICULAR APPARATUS: OPERATIONAL CHARACTERISTICS

OOCYTE-SOMATIC CELL INTERACTIONS

in premature depletion of the ovarian follicle reserve [71]. Therefore, this GC-derived hormone has been widely used as a biomarker of ovarian reserve and a reliable predictor of responsiveness to fertility induction therapy [72]. Further clinical studies revealed that AMH can be used to evaluate the ovarian reserve after chemotherapy or radiotherapy and predict the age at the start of menopause [73].

INTRAOVARIAN CELL-CELL COMMUNICATION After being recruited from the dormant primordial pool (gonadotropin-independent stage), most follicles develop into the FSH-dependent early antral stage [74]. In mono-ovular species, a dominant follicle in the cohort is selected and develops to the later antral stage when FSH is elevated [75]. While follicular dominance is highly dependent on the endocrine control that determines the cell fate (atresia) of the subordinate follicles, intraovarian interactions are also involved in this regulatory process [75]. In a mature antral follicle, coordinate communication between the oocyte and its supporting follicle cells (cumulus/granulosa and theca/stroma cells) relies heavily on functional gap junctions. Gap junctions are membrane channels that connect two neighboring cells. These channels have a particular selectivity mainly based on molecular size and allow the transfer of small molecules G and FSHB-211G> T polymorphisms in adult [71] and pubertal [74] women on FSH and other reproductive hormone levels, and suggested additive effects between the different genotypes. About 20% of people are carriers of the alleles associated with lower serum FSH levels/reduced FSHR expression or activity, possibly less favorable for reproduction. Larger prospective studies are needed to find out whether stratification of infertile patients according to their FSHR/FSHB genotypes improves clinical efficacy of FSH treatment.

CONCLUSION Concerning mutations in the gonadotropin subunit genes, the following conclusions can be drawn: • No germline mutations of the CGA are known, apparently because of the dramatic phenotypic effects they would have, with possible embryo lethality.

• Only eight inactive LHB mutations have been described, with normal prenatal sex differentiation but arrested pubertal development in men, and hypogonadotropic anovulatory infertility in women. • A common polymorphism exists in the LHB subunit. It affects the bioactivity of the hormone and has multiple mild phenotypic effects, including borderline compromising effects on female fertility. Its effect on reproductive functions may be strongly dependent on the ethnic background of an individual. • FSHB mutations in women cause disturbances in pubertal development and arrest follicular maturation. • FSHB and FSHR polymorphisms have limited impact on female reproduction, but certain FSHB/FSHR haplotype combinations may be more significant in affecting a woman’s FSH levels and response to FSH treatment upon IVF, offering thus a pharmacogenetic opportunity of individualized treatment. The following conclusions can be made about gonadotropin receptor mutations and polymorphisms: • Constitutively activating LHCGR mutations give rise to early-onset gonadotropin-independent precocious puberty in boys, but no female phenotype has been identified. • Two activating FSHR mutations have been described in men, but none women, and studies on candidate syndromes (e.g., familial twinning) have yielded negative results. • Some FSHR mutations induce relaxed ligand specificity toward hCG and account for pregnancyassociated ovarian hyperstimulation syndrome. • Inactivating LHCGR mutation causes a relatively mild phenotype in women: anovulatory infertility without apparent disturbance in pubertal development. The male phenotype ranges, depending on completeness of the receptor inactivation, from complete XY, DSD to mild undermasculinization with hypospadias and micropenis. • Inactivating FSHR mutations cause in women, depending on completeness of the loss of function, phenotypes ranging from FSH-responsive secondary to treatment-resistant hypergonadotropic primary amenorrhea with total arrest of follicular maturation, including arrested pubertal maturation. Despite their rarity, the gonadotropin and gonadotropin receptor mutations have been informative, because they have clarified the molecular pathogenesis of conditions associated with disturbed gonadotropin secretion and action. They have also clarified some poorly known or controversial issues in the physiology of gonadotropin function. This new information will make it possible to design more rational treatment regimens for gonadotropin therapy in general, and more specifically, for the cases where a genuine mutation in the genes involved has been

I. THE OVARIAN FOLLICULAR APPARATUS: OPERATIONAL CHARACTERISTICS

REFERENCES

discovered. Some of the common polymorphisms discovered in the genes of gonadotropins and their receptors may prove to be important contributing factors, in conjunction with specific genetic backgrounds, in the pathogenesis of certain reproductive disturbances, and may offer the opportunity to individualize hormonal therapies (pharmacogenetics). The structure-function relations of gonadotropin action at the three-dimensional level of molecule interaction will enable the design of small molecular form gonadotropin agonists and antagonists for future therapeutic applications.

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I. THE OVARIAN FOLLICULAR APPARATUS: OPERATIONAL CHARACTERISTICS

C H A P T E R

9 Environmentally Induced Epigenetic Transgenerational Inheritance of Ovarian Disease H.J. Kimbel, E.E. Nilsson, M.K. Skinner Abbreviations PCOS POI

polycystic ovarian syndrome primary ovary insufficiency

INTRODUCTION Epigenetics is defined as molecular factors and processes around the DNA that regulate genome activity independent of DNA sequence, and are mitotically or meiotically stable [1]. Epigenetic mechanisms can mediate changes in an organism’s phenotype that occur in response to environmental factors. Over several generations, shifts in the phenotype may occur in an organism’s lineage, even if the organism affected was not directly exposed to the toxicants. Ancestral exposure to a variety of environmental factors, toxicants, or drugs, has been shown to promote changes in gene expression and disease frequency [2]. This concept is known as multigenerational or transgenerational epigenetic inheritance. Transgenerational phenomena are inherited changes that do not involve continued direct exposure to an environmental factor, while multigenerational inheritance involves some type of direct exposures to the animal, developing embryo, or developing gametes within a developing embryo that then impact the different generations. Transgenerational epigenetic inheritance requires that epigenetic information or epigenetic changes are present in germ cells (i.e., sperm or eggs), as it is through germ cells that inheritance occurs. When conducting transgenerational epigenetic research, it is imperative that one evaluates the first generations that were not directly exposed to the toxicant and compare them to similarly bred vehicle control animals. Therefore, any abnormalities observed are known to have been transgenerationally inherited, rather than

The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00009-1

being due to epigenetic changes induced by direct toxicant exposure. There are two different exposure periods commonly investigated when evaluating transgenerational epigenetics in model species. One approach is to treat the F0 generation female during gestation. When using the rat model, female rats have been treated during pregnancy from embryonic developmental days 8 to 15 with a toxicant or vehicle control [3]. This is during the period of gonadal sex determination, at which time extensive epigenetic reprogramming occurs in the developing germ cells [4,5]. Exposure to environmental factors during this sensitive period may cause changes to the epigenome of the germ cells. Treatments administered at this time directly expose the developing pups (F1 generation) to the treatment, and the developing gametes within those pups, which will become the F2 generation. Thus, a transgenerational effect can only be seen in the F3 generation animals and beyond because those animals would not be directly exposed to any toxicants (see Fig. 1). The second approach used for examining transgenerational epigenetic changes is to expose an F0 generation male or nonpregnant female animal to a treatment and evaluate the F2 generation. This directly exposes the gametes within the F0 rats that will be used to form the F1 generation. Thus, any effects seen in the F2 generation can be considered transgenerationally inherited changes, since F2 generation animals were not directly exposed to environmental factors [6]. Ovarian disease affects a wide range of individuals throughout the world. The prevalence of ovarian disease varies by age and by population evaluated. The two most common types of ovarian diseases are polycystic ovarian syndrome (PCOS) and primary ovarian insufficiency (POI). PCOS affects 12%–21% of women of reproductive

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© 2019 Elsevier Inc. All rights reserved.

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9. ENVIRONMENTALLY INDUCED EPIGENETIC TRANSGENERATIONAL INHERITANCE OF OVARIAN DISEASE

Endocrine disruptor exposed gestating mother (sex determination period)

F0

F1

F2 germline

FIG. 1 Rat lineage example showing an F0 generation pregnant animal exposed to a toxicant, and illustrating that the multigenerational F1 and F2 generations were also directly exposed, while the transgenerational F3 generation had no direct exposure. Modified from Skinner MK. What is an epigenetic transgenerational phenotype? F3 or F2. Reprod Toxicol 2008;25(1):2–6.

age, depending on diagnostic criteria and the population assessed [7]. The onset of PCOS can be caused by genetic factors or environmental factors. PCOS in humans is characterized by a combination of different ovarian abnormalities. The prevalence of specific symptoms varies with age and with population. Three common symptoms are used for diagnosing the syndrome. An individual must have two of the three following conditions to be considered to have PCOS: Oligo/anovulation, hyperandrogenism, or polycystic ovaries on an ultrasound [8]. From these criteria, in humans, note that polycystic ovaries are not always present in people with PCOS. This poses an issue when evaluating nonhuman animals for PCOS-like conditions, as the only human PCOS criterion usually evaluated in animal models is the presence of polycystic ovaries. Primary ovary insufficiency is a condition characterized by follicular loss within the ovaries and menopause before the age of 40 in humans [9]. There are two mechanisms through which POI occurs: through follicle depletion or through follicle dysfunction [10]. Follicle depletion is a state in which there are very few follicles that remain

in the ovary. This can be either genetic, caused by an autoimmune disorder, or due to accelerated follicle expenditure [11]. Follicle dysfunction occurs when the follicles within the ovary remain but do not develop or ovulate. This can be due to exposure to endocrine disruptors or due to abnormalities, such as a mutation in the FSH receptor [12]. In the rat and mouse model, an animal with ovaries that are polycystic compared to the control indicates the presence of a PCOS-like condition. An animal with a significant reduction of primordial or developing follicles compared to controls indicates that the animal may have a condition similar to POI. Transgenerational epigenetic effects caused by different factors, such as pharmaceutical agents or environmental pollutants, can be evaluated both in terms of molecular changes induced within the germline epigenome of descendants and in characterizing observable phenotypic changes in descendants. When evaluating ovarian tissues, the number of primordial follicles, developing preantral follicles, and antral follicles may be counted. A low number of primordial or preantral

I. THE OVARIAN FOLLICULAR APPARATUS: OPERATIONAL CHARACTERISTICS

TRANSGENERATIONAL OVARIAN EFFECTS

follicles as compared to control animals often indicate disease. The number of cysts in each ovary is counted as well. If this number is high compared to controls that is likely indicative of abnormalities similar to PCOS. The signs that arise in rats and mice do not necessarily indicate the presence of the exactly equivalent human conditions of PCOS and POI, but instead these data can be used to better understand the underlying causes of similar ovarian diseases in rodents, and these findings can then be tested and correlated in human populations.

THE EPIGENOME Epigenetic processes are the molecular mechanism that organisms use to change gene expression in response to changes in their environment. DNA methylation was the first epigenetic process to be described [13]. DNA methylation occurs when a methyl group attaches to a cytosine molecule that is next to a guanine on the DNA strand. Following the discovery of DNA methylation, it was also discovered that histone modifications in the nucleosomes play a role in gene expression. Histone modifications can influence gene expression independent of DNA sequence. Chromatin structure can also modify gene expression. Noncoding RNA also play a role as an epigenetic factor [2]. Differential DNA methylated regions (DMRs) identify the presence of altered DNA methylation termed epimutations in the genome, compared to unaffected control animals. If DMRs are present in sperm or eggs, then epimutations may be passed through the germ line. DMRs are associated with transgenerational inheritance of disease susceptibility. Histone modification, ncRNA, and other epigenetic markers also likely play important roles in epigenetic transgenerational inheritance [14,15].

TRANSGENERATIONAL OVARIAN EFFECTS Several environmental toxicants have been studied to investigate potential transgenerational epigenetic effects they may have on the ovary (Table 1A and B). Toxicants studied in rats include vinclozolin [3], pesticides (permethrin and DEET) [3], plastic compounds [bisphenolA (BPA), dibutyl phthalate (DBP), and di(2-ethylhexyl) phthalate (DEHP)] [3], jet fuel [3], dichlorodiphenyltrichloroethane (DDT) [16], and methoxychlor [17]. In studies using rats as a model species, the animals were treated with doses generally equivalent to that of 1%–5% of the LD 50 (lethal dose for 50% of those treated) during the period of gonadal sex determination. At these doses, there was little overt toxicity in the directly exposed animals. These were not risk assessment studies, but rather

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pharmacological studies assessing the long-term epigenetic and transgenerational alteration effects that develop for each of the toxicants reviewed [3].

Vinclozolin Vinclozolin is a common agricultural fungicide that is used on fruits and vegetables, specifically wine grapes in the United States and is a known antiandrogen [19]. Vinclozolin is still used in some areas of the United States but its use has been decreasing since 2000, with a dramatic decrease in use from 2006 to today [20]. Following exposure of F0 generation gestating female rats to vinclozolin, the directly exposed F1 generation females aged to 1 year showed a significant increase in small ovarian cysts compared to controls. These F1 generation ovaries also had a significant decrease in primordial and developing follicles, reflecting a reduction in the total oocyte pool. Such decreases in the primordial follicle pool size may lead to lower fertility. In the transgenerational F3 generation of vinclozolin lineage animals the ovaries showed an increased incidence of both small and large ovarian cysts, as well as a significant decrease in follicle numbers [3] (see Table 1A and B). This F3 generation phenotype identifies epigenetic transgenerational inheritance of ovarian disease. Granulosa cells isolated from antral follicles of the transgenerational F3 generation vinclozolin lineage ovaries showed epigenetic differences compared to controls with 43 DMRs in the gene promoter areas, which are correlated with the environmentally induced transgenerational ovarian disease onset [3]. Genes previously shown to be associated with ovarian disease also had altered expression in these transgenerational granulosa cells.

Pesticides: Permethrin and DEET Permethrin is an insecticide used to treat scabies and lice when applied topically, and which is also used on cotton, wheat, and corn crops [21]. DEET (N,N-diethylmeta-toluamide) is the most common active ingredient in insect repellents, and is commonly applied topically to the skin [22]. Following exposure of F0 generation gestating female rats to a mixture of permethrin and DEET, the directly exposed F1 generation females showed a decrease in primordial follicles, indicating a lower follicle pool size. The transgenerational F3 generation rats also experienced a decrease in primordial follicle numbers, and an increase in both small and large cyst frequency as compared to F3 generation vehicle control lineage rats [3] (see Table 1A and B). This demonstrates a transgenerational effect of permethrin and DEET on ovarian disease.

I. THE OVARIAN FOLLICULAR APPARATUS: OPERATIONAL CHARACTERISTICS

152 TABLE 1

9. ENVIRONMENTALLY INDUCED EPIGENETIC TRANSGENERATIONAL INHERITANCE OF OVARIAN DISEASE

Ovarian Disease in Treated Lineage F1 (A) and Transgenerational F3 (B) Animals

Treatment

Primordial follicle decrease

Small cyst increase

Large cyst increase

(A)

F1 generation

Vinclozolin

+

+




Pesticides

+







DDT




+

+

Methoxychlor

+







Plastics

+

+

+

Dioxin

+







Jet Fuel

+

+




DEHP (mice)

+

n/a

n/a

Diseases of F1-treated lineage animals as compared to F1 control lineage animals. A plus sign (+) indicates that the abnormality was present [3,16–18]. (B)

F3 generation

Vinclozolin

+

+

+

Pesticides

+

+

+

DDT




+

+

Methoxychlor




+

+

Plastics

+

+

+

Dioxin

+

+

+

Jet Fuel

+

+

+

DEHP (mice)

+

n/a

n/a

Diseases of F3-treated lineage animals as compared to F3 control lineage animals. This shows a transgenerational effect [3,16–18].

Pesticide: DDT DDT is an environmentally persistent insecticide that was heavily used in the United States until its use was banned in 1972. Exposure to DDT may occur when consuming meat, fish, or dairy products. DDT is metabolized into dichlorodiphenyldichloroethylene (DDE) in the body and is stored in fatty tissues [23]. Following the treatment of pregnant F0 generation rats with DDT, the F1 generation females did not have polycystic ovaries or follicular insufficiencies. However, the transgenerational F3 generation DDT-lineage rats had an increase in incidence of small and large ovarian cysts compared to controls. Sixty percent of the F3 generation DDT lineage had polycystic ovaries compared to none in the F3 control lineage animals [16] (see Table 1A and B).

Methoxychlor does not build up in the food chain as do some toxicants, and it was developed as a replacement for DDT. However, research has shown that animals exposed directly to methoxychlor in food or water often experience an increased risk of ovarian and uterine disease, as well as infertility due to methoxychlor’s endocrine disrupting activity [24]. In rat studies in which F0 generation gestating females were exposed to methoxychlor, the following F1 generation rats did not experience a significant change in primordial follicle pool size, ovarian cysts, or overall ovarian disease as compared to the control lineage animals. The F3 generation methoxychlor lineage animals showed an increase in polycystic ovary incidence. This indicates that the rats showed symptoms of PCOS transgenerationally following ancestral exposure to methoxychlor [17] (see Table 1A and B).

Plastic Derived Compounds: BPA, DBP, and DEHP BPA, DBP, and DEHP are components of plastics that are environmental toxicants that humans are exposed to on a regular basis. BPA is a plastic component often used to make water bottles, medical equipment, and DVDs. BPA is also used to make epoxy resins, which can be used to coat pipes or metal soda cans. DBP is a common plasticizer that is used in adhesives and printer ink. It is used in nail polish and children’s toys, but its use in children’s toys has since been banned in the United States [25]. DEHP is used in the making of products containing polyvinylchloride (PVC) and is classified as a high production volume chemical [26]. After F0 generation gestating female exposure to a mixture of BPA, DBP, and DEHP, the F1 generation females were found to have fewer primordial follicles, and an increase in small and large cysts. In the F3 generation, there also was a transgenerational decrease in primordial follicles and an increase in small and large cysts [3] (see Table 1A and B). In a mouse study investigating BPA alone, after F0 generation pregnant animals were exposed it was found that the F3 generation had ovaries with altered expression of hormone receptor, steroidogenic, apoptotic, and antioxidant genes [27]. In a mouse study investigating DEHP alone, following exposure of the pregnant F0 generation animals, DEHP was found to cause alterations in the reproductive track, including reduced primordial follicle reserves, and reduced oocyte number and viability in the F2 and F3 generations of female mice. These changes were most pronounced when analyzing the F2 and transgenerational F3 generations [18] (see Table 1A and B).

Pesticide: Methoxychlor

Dioxin

Methoxychlor is pesticide used on agricultural crops and in animal feed. It is also known as DMDT.

Dioxin is an organic pollutant that is a byproduct of manufacturing processes. Some of these processes include

I. THE OVARIAN FOLLICULAR APPARATUS: OPERATIONAL CHARACTERISTICS

REFERENCES

paper bleaching, herbicide/pesticide manufacture, and solid/hospital waste incineration. It is a reproductive toxicant and can alter hormone levels and causes developmental abnormalities or cancer when direct exposure occurs. Exposure to dioxin is usually through foods such as meat, dairy, or fish [28]. Following the treatment of F0 generation gestating female rats with dioxin the F1 generation exposure lineage was found to have a significant decrease in primordial follicle numbers. The F3 generation animals also experienced a significant transgenerational decrease in primordial follicle pool size, as well as a significant increase in the incidence of small cysts [3] (see Table 1A and B).

Jet fuel: Hydrocarbon (JP8) JP-8 is a hydrocarbon used as jet fuel for aircraft. It has been used as a vehicle for pesticides, insecticides, and herbicides, and JP-8 is commonly used for dust control on gravel roads [29]. After F0 generation gestating female rats were administered a jet fuel treatment, the directly exposed F1 generation animals experienced a decrease in primordial follicle numbers, as well as an increase in small ovarian cysts. This indicates a lower follicle pool size and signs of PCOS. The transgenerational F3 generation experienced a decrease in primordial follicle pool size, as well as an increase in both small and large cyst incidence. This indicates a transgenerational decrease in oocytes and an increase in hallmarks of PCOS [3] (see Table 1A and B).

CONCLUSION The results from the previously mentioned studies provide valuable insight into the mechanisms of ovarian disease and show how environmentally induced epigenetic changes can be inherited and lead to an increased incidence of ovarian diseases. The expression of rodent ovarian disease can be different from that in humans, but the basic mechanisms involved are anticipated to be similar. It is interesting to note that the incidence of human ovarian diseases is now as high as 20% [30], and ancestral exposures to environmental toxicants must be considered as one underlying cause. Observations suggest that exposure to different environmental factors has the potential to alter the epigenome of an individual and promote ovarian disease transgenerationally in subsequent generations. This suggests that ancestral environmental factors can contribute to the etiology of ovarian disease cases in the current human population. The exposures to environmental stressors experienced by one’s great-grandmother during gestation could have lasting transgenerational effects in one’s

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own life and be inherited in subsequent generations. The high rate of ovarian disease will in part be due to such exposures and inherited transgenerationally. Further investigation of the molecular mechanisms involved will help elucidate the etiology of ovarian disease.

References [1] Skinner MK. Environmental epigenetic transgenerational inheritance and somatic epigenetic mitotic stability. Epigenetics 2011; 6(7):838–42. [2] Skinner MK. Endocrine disruptor induction of epigenetic transgenerational inheritance of disease. Mol Cell Endocrinol 2014;398 (1–2):4–12. [3] Nilsson E, Larsen G, Manikkam M, Guerrero-Bosagna C, Savenkova MI, Skinner MK. Environmentally induced epigenetic transgenerational inheritance of ovarian disease. PLoS One 2012;7(5):e36129. [4] Messerschmidt DM, Knowles BB, Solter D. DNA methylation dynamics during epigenetic reprogramming in the germline and preimplantation embryos. Genes Dev 2014;28(8):812–28. [5] Reik W, Dean W, Walter J. Epigenetic reprogramming in mammalian development. Science 2001;293(5532):1089–93. [6] Skinner MK. What is an epigenetic transgenerational phenotype? F3 or F2. Reprod Toxicol 2008;25(1):2–6. [7] March WA, Moore VM, Willson KJ, Phillips DIW, Norman RJ, Davies MJ. The prevalence of polycystic ovary syndrome in a community sample assessed under contrasting diagnostic criteria. Hum Reprod 2010;25(2):544–51. [8] Boyle J, Teede HJ. Polycystic ovary syndrome—an update. Aust Fam Physician 2012;41(10):752–6. [9] Laven JS. Primary ovarian insufficiency. Semin Reprod Med 2016;34(4):230–4. [10] Nelson LM, Anasti J, Flack M. Premature ovarian failure. In: Adashi EY, Rock JA, Rosenwaks Z, editors. Reproductive endocrinology, surgery, and technology. Philadelphia: Lippincott–Raven; 1996. [11] Nelson LM. Clinical practice. Primary ovarian insufficiency. N Engl J Med 2009;360(6):606–14. [12] Aittomaki K, Herva R, Stenman UH, Juntunen K, Ylostalo P, Hovatta O, et al. Clinical features of primary ovarian failure caused by a point mutation in the follicle-stimulating hormone receptor gene. J Clin Endocrinol Metab 1996;81(10):3722–6. [13] Holliday R, Pugh JE. DNA modification mechanisms and gene activity during development. Science 1975;187(4173):226–32. [14] Skinner MK, Guerrero-Bosagna C, Haque MM. Environmentally induced epigenetic transgenerational inheritance of sperm epimutations promote genetic mutations. Epigenetics 2015;10(8):762–71. [15] Nilsson EE, Skinner MK. Environmentally induced epigenetic transgenerational inheritance of disease susceptibility. Transl Res 2015;165:12–7. [16] Skinner MK, Manikkam M, Tracey R, Guerrero-Bosagna C, Haque MM, Nilsson EE. Ancestral dichlorodiphenyltrichloroethane (DDT) exposure promotes epigenetic transgenerational inheritance of obesity. BMC Med 2013;11:228. [17] Manikkam M, Haque MM, Guerrero-Bosagna C, Nilsson EE, Skinner MK. Pesticide methoxychlor promotes the epigenetic transgenerational inheritance of adult-onset disease through the female germline. PLoS One 2014;9(7)e102091. [18] Pocar P, Fiandanese N, Berrini A, Secchi C, Borromeo V. Maternal exposure to di(2-ethylhexyl)phthalate (DEHP) promotes the transgenerational inheritance of adult-onset reproductive dysfunctions through the female germline in mice. Toxicol Appl Pharmacol 2017;322:113–21.

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[19] Anway MD, Leathers C, Skinner MK. Endocrine disruptor vinclozolin induced epigenetic transgenerational adult-onset disease. Endocrinology 2006;147(12):5515–23. [20] United States GS. Estimated annual agricultural pesticide use 2014. updated January 17, 2017. Available from: http://water.usgs.gov/ nawqa/pnsp/usage/maps/show_map.php?year¼2014& map¼VINCLOZOLIN&hilo¼L&disp¼Vinclozolin. [21] Li W, Morgan MK, Graham SE, Starr JM. Measurement of pyrethroids and their environmental degradation products in fresh fruits and vegetables using a modification of the quick easy cheap effective rugged safe (QuEChERS) method. Talanta 2016;151:42–50. [22] Osimitz TG, Grothaus RH. The present safety assessment of DEET. J Am Mosq Control Assoc 1995;11(2 Pt 2):274–8. [23] CDC. Dichlorodiphenyltrichloroethane (DDT) 2016 [updated December 23, 2016. Available from: https://www.cdc.gov/ biomonitoring/ddt_factsheet.html. [24] ATSDR. 2002. Toxic substances portal—methoxychlor. Atlanta, GA: Department of Health and Human Services, Public Health Service. Available from: https://www.atsdr.cdc.gov/toxfaqs/tf. asp?id¼777&tid¼151. [25] Consumer Product Safety Improvement Act of 2008 Public Law 110-314 (2008). [26] Lorz PM, Towae FK, Enke W, J€ackh R, Bhargava N, Hillesheim W. Phthalic acid and derivatives. In: Ullmann’s encyclopedia of industrial chemistry. Weinheim: Wiley-VCH; 2002. [27] Berger A, Ziv-Gal A, Cudiamat J, Wang W, Zhou C, Flaws JA. The effects of in utero bisphenol a exposure on the ovaries in multiple generations of mice. Reprod Toxicol 2016;60:39–52. [28] WHO. Dioxins and their effects on human health [updated October 2016. Available from: http://www.who.int/mediacentre/ factsheets/fs225/en/.

[29] Ritchie G, Still K, Rossi 3rd J, Bekkedal M, Bobb A, Arfsten D. Biological and health effects of exposure to kerosene-based jet fuels and performance additives. J Toxicol Environ Health B Crit Rev 2003;6(4):357–451. [30] Sirmans SM, Pate KA. Epidemiology, diagnosis, and management of polycystic ovary syndrome. Clin Epidemiol 2013;6:1–13.

Glossary DNA methylation An epigenetic process in which a methyl group attaches to a cytosine molecule that is next to a guanine on the DNA strand. Epigenetics Molecular factors and processes around DNA that regulate genome activity independent of DNA sequence, and which are mitotically or meiotically stable. Polycystic ovarian syndrome (PCOS) An individual having oligo/anovulation, hyperandrogenism, or polycystic ovaries on an ultrasound. In humans, PCOS is characterized as an individual having two of the three above conditions. Primary ovary insufficiency (POI) A condition characterized by marked follicular loss within the ovaries and menopause before the age of 40 in humans. Transgenerational epigenetic inheritance Germline (sperm or egg) transmission of epigenetic information between generations in the absence of any direct exposures or genetic manipulations.

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10 Mammalian Oogenesis: The Fragile Foundation of the Next Generation John J. Bromfield, Rachel L. Piersanti INTRODUCTION Development of the mammalian oocyte requires a series of orchestrated events over a long period of time to facilitate the formation of a single competent haploid cell. The process of meiosis is a unique series of cellular divisions which occurs in both male and female gametes. The goal of meiosis is to reduce the number of chromosomes from a diploid configuration to a haploid number. This process ensures that upon fertilization the newly formed zygote has half of its genetic makeup from the sire and half from the dam ensuring a unique genetic makeup of the new offspring. The competent development of the oocyte is integral to the processes of fertilization and healthy diploid embryo development. Oocyte development requires cytoplasmic maturation of the cell, to facilitate the cellular requirements of the early embryo during early cleavage events; nuclear maturation, to ensure the development of a unique haploid cell which is arrested just prior to completion of meiosis and is prepared for fertilization; and epigenetic regulation of the oocyte genome, required for appropriate gene expression throughout development in a heritable manner. Only when these processes are completed correctly can the oocytes undergo fertilization and develop into an embryo and subsequently a healthy progeny. Any mistakes during these processes can have significant negative consequences to completing oocyte maturation or early embryo development.

membranes of the yolk-sac (mesoderm) at embryonic day 7 [1]. These PGCs then migrate through the hindgut and colonize the genital ridge which will become the embryonic gonad. During migration through the hindgut, PGCs undergo mitotic cellular proliferation and become oogonia. In the mouse, the process of mitosis ends at approximately embryonic day 13 and the population of finite oocytes is established [2]. The newly formed oocyte population can now begin the cellular division of meiosis. However, it is important to note that oocytes at this stage of development arrest at the dictyate stage of prophase I. It is well documented that mammalian oocytes are a finite population of cells, as such the number of oocytes formed at this stage of development will constitute the ovarian reserve for the life of the animal. This paradigm has been challenged however, to suggest an ability of oocytes to be replenished by an oogonial stem cell population [3,4]. Nonetheless, these studies remain controversial and the existence or physiological role of such cells remains in question [5]. Once the ovarian reserve is established, oocytes must undergo a series of developmental milestones (listed above) to become developmentally competent cells which can be fertilized and give rise to healthy embryos and offspring. This process can take years following the initial development of the oocyte, progression through puberty, and finally completion of the meiotic cell cycle only after fertilization.

CYTOPLASMIC MATURATION EMBRYONIC DEVELOPMENT OF THE OVARIAN RESERVE The process of meiosis in the female begins during fetal development. Primordial germ cells (PGCs) are first discernable in the female mouse in the extraembryonic

The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00010-8

Cytoplasmic maturation is the process by which oocytes prepare their internal structure and contents to facilitate fertilization and early embryonic development. Fertilization itself initiates a number of molecular events within the cytoplasm of the oocyte required for progression into embryonic development. As such the oocyte

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cytoplasm must be competent to facilitate these early postfertilization events to allow embryonic development. The newly formed embryonic genome does not become active until a number of cellular cleavages have occurred [6], as such the early embryo is dependent on the maternal supplies laid down in the oocyte during development for cellular activity and division.

Maternal Effect Genes During the process of development, oocytes accumulate maternal effect factors such as proteins, lipids, mitochondria, and RNA. All of these factors are critical to the early cellular divisions of the newly formed zygote as the sperm carries little more than DNA into the process of fertilization. Maternal effect genes are synthesized and stored during oocyte growth. Upon synthesis these RNA molecules, which code for key molecules required for oocyte to embryo transition, remain free of polyadenylation and as such are not transcribed [7]. Only upon polyadenylation of the 30 region of the RNA molecules do these transcripts which become subject to translation [8]. While Drosophila and Xenopus have proven to be valuable models in identifying maternal effect genes in oocytes; in rodents (and other mammals) little is currently known about the identity of these maternal effect genes [9,10]. The maternal effect gene, zygotic arrest-1 (Zar1) is an oocyte specific factor critical in facilitating the transition from a 1-cell zygote to a 2-cell embryo. When Zar1 is genetically deleted from female mice, embryonic development does not progress beyond the 1-cell zygote stage [11]. Beyond the mouse a role for ZAR1 has been identified in cattle, pigs, and humans [12,13]. Another maternal effect gene to be identified in mammals is NOD-like receptor family pyrin domain containing 5 (Nlrp5; formerly known as Mater) [14]. Expression of Nlrp5 RNA is oocyte specific and is not detected once the embryonic genome is activated. Indeed, when Nlrp5 is genetically deleted from female mice, embryos cannot develop beyond the 2-cell stage when embryonic genome activation occurs. Conversely, males which have had Nlrp5 deleted do not show any disruption to fertility, suggesting maternally derived Nlrp5 is critical for embryo development. The importance of Nlrp5 as a maternal effect gene has also been described in sheep, cattle, and pig, while a human homologue has also been characterized [13,15–18].

Cytoplasmic Reorganization As an oocyte develops it undergoes structural and functional reorganization of its intracellular machinery. These functional and structural changes occur in a precise spatiotemporal manner to facilitate the specific

requirements of the oocyte as it progresses through development. Cellular organelles are moved around the oocyte cytoplasm to alter their function dependent on the stage of oocyte development. Mitochondria, the power generators of the oocyte, are solely maternally derived. Mitochondria are critical during development as a source of oxidative phosphorylation to provide the oocyte with the required ATP for cellular activity. Mitochondria are crucial to the function of the oocyte due to its inability to metabolize glucose efficiently [19]. The number of mitochondria present in the oocyte is variable between species; mice have approximately 92,000 mitochondria, a while human oocytes contain between 30,000 and 1,000,000 mitochondria [20–22]. This number is thought to change during oocyte development with mitochondrial copy number being highest during prophase arrest and subsequently decreasing during final maturation to the MII stage [23]. While the total number of mitochondria is important in the energy output capacity of the oocyte, the cellular distribution of these organelles also seems to affect their functionality. In the mouse, mitochondria are dispersed throughout the oocyte cytoplasm during prophase I arrest, and at the resumption of the cell-cycle mitochondria begin to aggregate around the newly formed meiotic spindle (MI and MII) [24]. It is believed that this aggregation of mitochondria at the meiotic machinery is deliberate and concentrates these power houses to the site of major energy consumption—spindle formation and chromosome segregation are high-energy processes [25]. Finally the distribution of mitochondria upon first polar body extrusion also appears asymmetrical, with the majority of mitochondria being positioned at the inner pole of the meiotic spindle [24]. This repositioning toward the interior of the oocyte ensures the maximal number of mitochondria are carried forward with the oocyte and not lost to the polar body. The endoplasmic reticulum (ER) is a membranous organelle involved in protein synthesis and transport (rough ER), and lipid and steroid (smooth ER) synthesis. While these organelles serve their traditional function in the oocyte, they also serve as a significant calcium (Ca2+) store for rapid intracellular signaling involved in the postfertilization reaction [26]. The release of ER stored Ca2+ is of critical importance to the timing of events following fertilization. The release of ER Ca2+ triggers the cortical reaction by the cortical granules preventing polyspermy, and initiating completion of meiosis and polar body extrusion [27]. Release of Ca2+ by the ER is initiated by the sperm associated factor phospholipase C-zeta (PLCζ) which increases oocyte inositol trisphosphate (IP3) which binds to its receptor on the ER membrane and facilitates Ca2+ efflux [26]. The spatial distribution of the ER changes as the oocyte progress through development with the ER dispersed throughout the cytoplasm of the oocyte during prophase I arrest. However, the

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distribution of the ER is located at the oocyte cortex at the MII arrest with an apparent absence of ER nearest the oocyte membrane proximal to the spindle [28]. This spatial redistribution of the ER correlates with the capacity of the oocyte to release Ca2+ in response to fertilization, when the ER is dispersed during prophase I arrest the capacity to release Ca2+ is reduced, while at MII when the ER is cortically located the capacity to secrete Ca2+ is greatest [29,30]. This situation fits perfectly with the Ca2+ requirements of the oocyte to facilitate Ca2+dependent postfertilization events. Stiffening of the cellular cortex is a hallmark of symmetrical mitotic cell division [31]. Surprisingly, the asymmetrical cell division in oocytes, which gives rise to a small polar body and large oocyte, occurs with a decrease in overall cortical stiffening and the greatest reduction occurs at the site of polar body extrusion [32]. The observed reduction in cortical stiffness occurs in parallel with an increase in cortical actin polymerization during oocyte development and as such is not a passive artifact of reduced cytoskeletal strength, but a targeted reduction thought to be critical to facilitate asymmetrical cell division [33]. Taken together these changes to the cytoplasm are critical in the developmental competence of the oocyte to become fertilized, activate the embryonic genome, and undergo embryonic development to result in healthy offspring [34–36].

NUCLEAR MATURATION Nuclear maturation is a mechanism by which oocytes reduce their genetic material by half to result in a haploid cell (n). The process of nuclear maturation itself occurs during the entire life of the oocyte, beginning during fetal developmental and is not completed until fertilization

(Fig. 1). After the completion of oogonial mitotic proliferation (described above), newly formed oocytes enter the meiotic cell cycle during fetal life. Oocytes enter the meiotic cell cycle and quickly arrest at prophase of the first meiotic cell cycle. Prophase I arrested oocytes will exist as primordial follicles until acquiring an activation signal to initiate growth; however this growth will focus on hypertrophy of the oocyte and cytoplasmic maturation (as above) while the oocyte remains arrested at prophase I until fully grown. These oocytes are referred to as germinal vesicle (GV) oocytes and are characterized by an intact nuclear membrane. Primordial follicles consist of a single layer of squamous (flattened) somatic cells termed granulosa cells, and a prophase arrested oocyte. Granulosa cells of the follicle are a critically important component of the follicle in regulating the nuclear maturation of oocytes as they continue to grow from the primordial stage to large ovulatory follicles. Upon activation, a primordial follicle transitions into a primary follicle which is characterized by a histological change in granulosa cells from squamous to cuboidal in shape. The interaction between granulosa cells and oocytes is integral in dictating the function of both granulosa and oocytes. As the follicle progresses in development, granulosa cells proliferate increasing the number of layers surrounding the now growing oocyte. The formation of an antral space in the granulosa cell population dictates the formation of an antral follicle containing a fully grown oocyte [37]. The transition from a preantral to antral follicle marks the ability of the oocyte to reinitiate the meiotic cell cycle; indeed, the capacity for oocytes to complete meiosis is dependent on oocyte growth [38]. It is important to note here that the finite population of follicles established at birth in the mammalian ovary will not all result in complete follicle growth and ovulation. The vast majority of follicles established at birth will undergo FIG. 1

Control of cell cycle during nuclear maturation. Activity of maturation promoting factor (MPF) is defined by the phosphorylation status of CDK1 (active, dephosphorylated—green; inactive, phosphorylated—red) and availability of CYB (degraded—gray; available— blue). CDK1 phosphorylation is regulated by the phosphatase CDC25 which is in turn regulated by the kinase WEE1.

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cell death (atresia) during any one of the follicle stages, resulting in a number of primordial follicles activated in a single wave to give rise to a single ovulatory follicle. Growth of the oocyte during preantral follicular development is coordinated with granulosa cell proliferation and is regulated by the communication between the two cell types. Growth differentiation factor 9 (GDF9) and bone morphogenic factor 15 (BMP15) are oocyte specific secreted factors which act on granulosa cells to coordinate oocyte growth and granulosa cell proliferation, maintaining orchestrated growth between the two cell types [39,40]. Genetic depletion of Gdf9 in mice results in a physical disconnect between oocytes and granulosa cells, giving rise to abnormally large oocytes that do not complete meiosis [41,42]. This communication between oocytes and granulosa cells appears to be bidirectional and requires negative feedback of growth mechanisms in the oocytes. The preovulatory surge of pituitary luteinizing hormone (LH) is the initiating factor which resumes meiotic cell cycle in the fully grown oocyte (Fig. 2). Granulosa cells in the preovulatory antral follicle increase expression

FIG. 2

Resumption of meiosis by LH. Engagement of the LH receptor on cumulus cells activates adenylate cyclase increasing intracellular cAMP. Elevated cAMP increases MAPK activity and increases PDE in the oocyte. PDE depletes intra-oocyte cAMP and dissociation of cumulus cells from the oocyte reduces cAMP transport to the oocyte. Reduced intra-oocyte cAMP activates CDC25 resulting in dephosphorylation of CDK1 promoting resumption of meiosis.

of LH receptors prior to ovulation [43] and it is believed that engagement of the LH receptor in granulosa cells is the initiating factor to resume meiosis in the oocyte. Meiotic cell cycle arrest during quiescence and follicle growth up until ovulation is maintained by elevated oocyte cyclic adenosine monophosphate (cAMP). The use of cAMP analogs is a useful tool to maintain cell cycle arrest in oocytes, while phosphodiesterase (PDE) inhibitors can be used to prevent the degradation of endogenous oocyte cAMP and maintain cell cycle arrest [44,45]. Cyclic guanosine monophosphate (cGMP) derived from the surrounding cumulus cells has also been proven to diffuse into the oocyte to maintain oocyte cAMP levels and to maintain cell cycle arrest [46]. Protein kinase A (PKA) is activated by cAMP, as such PKA is a potential mediator of cell cycle activation in the oocyte by regulating important cell cycle mediators (see below). It is postulated that LH binding to the LH receptor on cumulus cells initiates physical dissociation of cumulus cells from the oocyte, resulting in decreased transport of cAMP into the oocyte [47]. Additionally, LH-mediated cell cycle resumption also requires the activation of MAPK1 in cumulus cells. It has been surmised that cumulus MAPK increases PDE activity in the oocyte resulting in decreased cAMP and subsequent meiotic resumption [48,49]. Upon removal of the cell cycle inhibitory signals, oocytes can continue to undergo meiotic cell division (Fig. 1). Similar to somatic cells undergoing mitosis, the oocyte relies on cyclin B (CYB) and cyclin-dependent kinase 1 (CDK1) [50,51]. In combination, these two proteins form a complex referred to as maturation promoting factor (MPF). During prophase arrest CDK1 protein is maintained in an inactive state by phosphorylation of the catalytic subunit by the kinases MYT1 and WEE1 [52–55]. The CDK1 protein is activated by dephosphorylation of the catalytic subunit by the phosphatase CDC25 [56,57]. The activity of CDC25 is regulated at the level of phosphorylation also; when inactive CDC25 is maintained in a phosphorylated state by PKA whose activity is maintained in the presence of high cAMP. Taken together, elevated cAMP maintains PKA activity, PKA inactivates CDC25 which is required to dephosphorylate CDK1 and facilitate cell cycle resumption. In addition to CDK1 activity, MPF is regulated by CYB degradation [58,59]. Oocyte CYB increases dramatically at GV breakdown (GVBD) to facilitate the onset of cell cycle resumption; however at completion of the first meiotic cell cycle CYB is rapidly degraded by ubiquitination and CDK1 is re-phosphorylated to initiate the second meiotic cell cycle arrest at metaphase (M) II. Meiosis is not completed until after fertilization, prior to which oocytes are arrested at M-phase of the second meiotic division around the time of ovulation. While cAMP is important in maintaining the first cell cycle arrest, cytoplasmic factors of the MII oocyte are critical

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in maintaining the second cell cycle arrest. The arresting factor was first elucidated when the cytoplasmic contents of an arrested MII oocyte were injected into the cytoplasm of a 2-cell embryo [60]. This transfer of cytoplasmic contents resulted in abrupt cell cycle arrest in the newly formed embryo. Studies in rodents to determine the identity of the cell cycle arrest factor have revealed MOS, MAPK1, and MAP2K1 as essential elements in cell cycle arrest at MII [61–63]. In the absence of these factors cell cycle arrest does not occur and oocytes progress through the second meiotic cleavage and undergo parthenogenetic activation [64].

EPIGENETIC REGULATION OF THE GENOME Epigenetic marks refer to alterations to the chromosome structure that affect gene expression without altering the DNA sequence in a heritable manner. Epigenetic reprogramming during oogenesis is essential for embryo and offspring development. This section focuses on new findings on epigenetic reprogramming during oogenesis and how those changes can be heritable to the following generations. In mammals, the main epigenetic regulatory mechanisms include DNA methylation, and posttranslational histone modifications, such as acetylation, phosphorylation, and methylation. These epigenetic modifications can be found at coding regions, promoters, and enhancers, and function as modulators of gene expression [65]. An important process during oogenesis is the establishment of genomic imprints. Imprinting is a procedure in which one of the parental alleles is marked, or imprinted; so only one allele will be preferentially or individually expressed. DNA methylation is the main mechanism for establishment of genomic imprints [66,67]. During two stages of mammalian development the epigenetic marks are erased and reestablished. The first time the epigenetic information is erased occurs right when the PGCs arrive at the genital ridge, as they undergo extensive DNA demethylation. The process happens by active demethylation when the proteins TET1 and TET2, oxidize 5-methylcytosine to 5-hydroxymethylcytosine [68] but also by passive demethylation via reduced expression of DNMT1, the enzyme responsible for maintaining methylation, which leads to a gradual dilution of the methylation patterns through DNA replication. Demethylation is also helped by reduction in expression of de novo DNA methyltransferases DNMT3A and DNMT3L; although the knockout of these two enzymes seems to have minor effects for oocyte development, the embryos lacking Dnmt3a or Dnmt3l fail to develop [69]. This demethylation reprograming in PGCs affects even the imprinted regions,

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but excludes intracisternal particle retrotransposon family (IAP), and several CpG islands within the genome [70,71]. The maintenance of some level of methylation in specific regions is one of the key evidences for epigenetic inherence and the study of fetal programing [72]. The degree of methylation erasure is greater in PGCs than in the preimplantation embryo, which is the second time point during development when the epigenetic marks will be erased [73]. It is believed that epigenetic reprograming in PGCs is important not only to establish transgenerational marks but mainly for appropriate development of the PGCs and future gametes. Another key event during this period is the reactivation of the X chromosome, which begins during the early stages of PGCs migration and it is completed in post-migratory PGCs [74]. After mitotic replication of the PGCs, somatic granulosa cells surround them and each oocyte is encapsulated in a primordial follicle. The oocyte remains unmethylated while in the primordial follicle and gradually becomes methylated during its growth [66]. There is no active transcription during oocyte maturation but there is activation and repression of specific mRNAs responsible for carrying out development until the embryonic genome is activated after fertilization. After fertilization the parental genomes undergo extensive demethylation at different rates for the maternal and paternal genomes but conserve methylation on imprinted genes and retrotransposons [67]. These mechanisms suggest that epigenetic marks obtained during oocyte development can be erased, but also that epigenetic inheritance can occur through non-erasure of methylation in specific regions of the genome. The activation of the embryonic genome is the process where the preimplantation embryo stops depending on the maternal transcripts from the oocyte and starts relying on its own transcription for development. Embryonic genome activation happens at different stages among different species, for example it occurs around the 4-cell stage in humans, and 8-cell stage in the bovine and rabbit [65]. As previously mentioned, posttranslational histone modifications are also important mechanisms for establishment of epigenetic marks. A specific group of posttranslationally modified histones is present during oocyte maturation, and in grown oocytes the histones H3 and H4 are abundant in their acetylated form. As the oocyte undergoes meiotic maturation, the chromatin suffers major deacetylation and remains that way to metaphase II [75]. The deletion of deacetylation regulatory enzymes has been studied and used to prove the importance of this posttranslational modifications for oocyte growth and development, deletion of the enzymes HDAC1 and HDAC2 lead to oocyte developmental arrest [76]. During embryo development there are differences in

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the distribution of histone marks between the parental genomes, however, although many studies were able to identify such differences, there is still no clear understanding of its function and how this epigenetic alteration impacts the regulation and control of gene expression [77]. Recently studies have been targeting intergenerational inheritance and fetal programming. Many of these studies focus on exposing the mother to specific diets or diseases during different periods of development, and evaluating the effects on the offspring. It has been reported that even if the mother is only exposed to a challenge for a short window of time before mating, it can have detrimental effects on offspring development [78]. This reinforces the idea that the environment can affect the oocyte during the final stages of development, when they are transcriptionally inactive and shows the importance of the epigenetic modifications during different stages of oogenesis and the potential to compromise the embryo quality. In summary, epigenetic marks are responsible for regulating gene expression without altering the DNA sequence. Epigenetic regulation is extremely important during oogenesis and embryo development, especially during the two windows of time where the epigenetic marks are erased and reestablished. During these specific time points there is potential for disruptions that may affect not only the oocyte, but also the offspring and the following generations.

CONCLUSIONS The process of oocyte maturation is unique. No other cell type grows for as long and may never see the completion of its cell cycle (meiosis is only completed after fertilization). Such a large cell that may exist in a quiescent state for such a long period of time (from birth to menopause) is truly a source of scientific curiosity. Many unanswered questions remain regarding the molecular events which govern the growth and development of such a unique cell. Future insights will infinitely help our basic understanding of mammalian reproductive physiology, human infertility, contraception, stem cells, agricultural production, and species preservation. Most recently researchers have been able to recapitulate the entire process of oocyte development in vitro from either PGCs or induced stem cells, fertilized these oocytes, generated offspring, and further recapitulated the events from those subsequent young [79,80]. These types of advancements in technology will allow a deeper understanding into the cellular and molecular queries governing oocyte development which remain to be elucidated.

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[66] Smallwood SA, Tomizawa S, Krueger F, Ruf N, Carli N, SegondsPichon A, et al. Dynamic CpG island methylation landscape in oocytes and preimplantation embryos. Nat Genet 2011;43(8):811–4. [67] Kobayashi H, Sakurai T, Imai M, Takahashi N, Fukuda A, Yayoi O, et al. Contribution of intragenic DNA methylation in mouse gametic DNA methylomes to establish oocyte-specific heritable marks. PLoS Genet 2012;8(1):e1002440. [68] Hackett JA, Sengupta R, Zylicz JJ, Murakami K, Lee C, Down TA, et al. Germline DNA demethylation dynamics and imprint erasure through 5-hydroxymethylcytosine. Science 2013;339(6118):448–52. [69] Bourc’his D, Xu GL, Lin CS, Bollman B, Bestor TH. Dnmt3L and the establishment of maternal genomic imprints. Science 2001; 294(5551):2536–9. [70] Lane N, Dean W, Erhardt S, Hajkova P, Surani A, Walter J, et al. Resistance of IAPs to methylation reprogramming may provide a mechanism for epigenetic inheritance in the mouse. Genesis 2003;35(2):88–93. [71] Hajkova P, Erhardt S, Lane N, Haaf T, El-Maarri O, Reik W, et al. Epigenetic reprogramming in mouse primordial germ cells. Mech Dev 2002;117(1–2):15–23. [72] Chong S, Whitelaw E. Epigenetic germline inheritance. Curr Opin Genet Dev 2004;14(6):692–6. [73] Guibert S, Forne T, Weber M. Global profiling of DNA methylation erasure in mouse primordial germ cells. Genome Res 2012;22(4):633–41.

[74] Sasaki H, Matsui Y. Epigenetic events in mammalian germ-cell development: reprogramming and beyond. Nat Rev Genet 2008;9(2):129–40. [75] Clarke HJ, Vieux KF. Epigenetic inheritance through the female germ-line: the known, the unknown, and the possible. Semin Cell Dev Biol 2015;43:106–16. [76] Ma P, Pan H, Montgomery RL, Olson EN, Schultz RM. Compensatory functions of histone deacetylase 1 (HDAC1) and HDAC2 regulate transcription and apoptosis during mouse oocyte development. Proc Natl Acad Sci U S A 2012;109(8):E481–9. [77] Canovas S, Ross PJ. Epigenetics in preimplantation mammalian development. Theriogenology 2016;86(1):69–79. [78] Wu G, Bersinger NA, Mueller MD, von Wolff M. Intrafollicular inflammatory cytokines but not steroid hormone concentrations are increased in naturally matured follicles of women with proven endometriosis. J Assist Reprod Genet 2017;34(3): 357–64. [79] Hayashi K, Ogushi S, Kurimoto K, Shimamoto S, Ohta H, Saitou M. Offspring from oocytes derived from in vitro primordial germ cell-like cells in mice. Science 2012;338(6109): 971–5. [80] Hikabe O, Hamazaki N, Nagamatsu G, Obata Y, Hirao Y, Hamada N, et al. Reconstitution in vitro of the entire cycle of the mouse female germ line. Nature 2016;539(7628):299–303.

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11 Regulation of Mammalian Oocyte Maturation Maria M. Viveiros, Rabindranath De La Fuente INTRODUCTION Mammalian oocyte maturation involves key developmental programs, which are essential for the production of a mature egg that is competent to undergo fertilization and support the initial stages of embryogenesis up to the blastocyst stage. This chapter addresses two crucial oocyte maturation processes, nuclear and epigenetic maturation. (1) Nuclear maturation includes the resumption and completion of the first meiotic division as well as the maintenance of a stable metaphase-II arrest. (2) Epigenetic maturation occurs during oocyte growth and results in crucial genomic modifications that regulate gene expression, nuclear architecture, and chromosome stability.

NUCLEAR MATURATION Meiosis is a unique cell division that occurs in gametes to reduce the number of chromosomes from a diploid (2 N) to haploid (N) number, such that upon fertilization a single copy of each maternal and paternal chromosome is transmitted to the embryo. In female mammals meiosis is initiated early during fetal development. Primordial germ cells (PGCs) are first discernible with the extraembryonic mesoderm of the mouse embryo at day 6.5 (E6.5), then migrate through the hindgut and dorsal mesentery to colonize the genital ridge [1,2]. Throughout their migration PGCs proliferate, however, around E12.5 mitosis ceases and the entire cell population enters prophase of the first meiotic division (meiosis-I) [3]. This change from mitotic proliferation into the first meiotic division defines the transition from oogonia to oocyte. Fetal oocytes then progress through pachytene and early diplotene stages necessary for meiotic recombination, but subsequently all arrest at the dictyate stage of prophase-I. Completion of the first meiotic division will occur only after the oocyte and the ovarian follicle have undergone extensive growth postnatally.

The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00011-X

In the perinatal ovary, following meiotic arrest, oocytes become surrounded by a single (squamous) layer of pregranulosa follicular cells and form primordial follicles. Therefore, at birth the ovaries contain a mammalian female’s full complement of oocytes, all within the large population of nongrowing primordial follicles [4]. Nevertheless, only a small fraction of the oocytes will ultimately be ovulated and have the potential to be fertilized. The majority of oocytes, together with the follicular somatic cells, will undergo atretic degeneration. Ovarian follicles that escape atresia and are recruited from the primordial pool become primary follicles. The oocyte begins an extensive growth phase during which the surrounding somatic cells (now called granulosa cells) become cuboidal and proliferative. Complex interactions between the oocyte and granulosa cells are essential for the development and function of both cell types. As follicles progress to the secondary stage, they are characterized by oocytes at mid-growth surrounded by multiple layers of granulosa cells. Follicles then develop to the antral stage containing fully grown oocytes within the follicular fluid-filled antrum. Two distinct granulosa cells are evident in antral follicles: (i) cumulus granulosa cells that directly surround the oocyte, and several outer layers of (ii) mural granulosa cells. Gap junction complexes connect all granulosa cells within the follicle as well the cumulus cells to the oocyte. The transition from preantral to antral stage is a critical juncture during which the oocyte acquires the capacity to resume meiosis [5]. Remarkably, mammalian oocytes can sustain prophase-I arrest for an extensive period (months or years, depending on the species) from the fetal stage until after puberty (sexual maturity), when during each reproductive cycle a cohort of ovarian follicles grow to the preovulatory stage. Fully grown oocytes within antral follicles normally resume meiosis in response to the preovulatory surge of luteinizing hormone (LH). Oocytes arrested at prophase-I are characterized by a large nucleus with a pronounced nucleolus. The oocyte nucleus is commonly referred to as the “germinal vesicle”

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(GV). Initial indicators of meiotic resumption include the breakdown of the nuclear membrane (“germinal vesicle breakdown,” GVB) along with the disappearance of the nucleolus. Upon GVB, the chromatin rapidly condenses into individual chromosomes, and a bipolar metaphase-I (MI) spindle is assembled. During the first meiotic division (meiosis-I), homologous chromosomes segregate and one set is extruded into the first polar body (Fig. 1). There is no intervening DNA synthesis and the chromosomes remain condensed. A second meiotic spindle is then assembled as oocytes enter metaphase-II (MII), where they normally arrest again (Fig. 1). The second meiotic division (meiosis-II) is only initiated upon fertilization and entails the segregation of sister chromatids, with one set extruded into a second polar body. In most mammals meiotic maturation to the MII stage is completed by the time of ovulation. Nevertheless, oocyte maturation and ovulation are not necessarily linked. Early studies revealed that spontaneous (LHindependent) maturation occurs when oocyte-cumulus cell complexes are removed from large antral follicles

and cultured in supporting medium [6,7], establishing that gonadotropin-mediated induction of ovulation is not essential to initiate nuclear maturation in the oocyte. Conversely, precocious induction of ovulation by gonadotropin treatment early in the estrous cycle of pigs results in the ovulation of immature primary (diploid) oocytes [8]. Notably, in contrast to other mammals, oocytes of dogs and foxes (Canidae) are ovulated at the GV stage and undergo meiotic maturation within the oviduct [9,10], further supporting that ovulation and oocyte maturation to MII are not strictly interdependent.

Key Cell Cycle Molecules Driving Nuclear Maturation Control of the resumption and progression of meiotic division in the oocyte is critical. Advancements in several areas of investigation have contributed to an increased understanding of the underlying mechanisms that regulate these processes. Similar to mitosis [11] in somatic

FIG. 1 Nuclear maturation of mouse oocytes. DNA is visualized by staining with propidium iodide (red) and anti-β-tubulin was used for labeling the spindle microtubules (green). Upper panel shows germinal vesicle (GV) stage oocyte with a condensed surrounded nucleolus (SN) configuration. Oocyte at metaphase I (MI) shows homologous chromosomes (red) aligned on the metaphase plate and attached to microtubules on a barrel-shaped spindle (green). Oocyte arrested at the metaphase (MII) stage with an extruded first polar body. Lower panel shows a phase contrast micrograph of the GV stage oocyte as well as corresponding chromosome configurations at metaphase I (MI) and metaphase (MII).

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cells, meiosis is regulated by oscillations in the activity of a key protein kinase, cyclin-dependent kinase 1 (CDK1/ p34cdc2) and its regulatory subunit cyclin B1. CDK1 activity was first described in Xenopus oocytes and initially referred to as “maturation promoting factor (MPF).” [12–14] Studies have since demonstrated that this kinase plays an essential role in regulating both meiotic resumption and progression. In mammalian oocytes, CDK1 is activated shortly before meiosis resumes and is necessary for GVB. Oocyte-conditional knockdown of cdk1 in mice leads to the failure of meiotic resumption and female infertility [15]. Following GVB, CDK1 activity normally increases and plateaus at MI, then decreases significantly during anaphase-I onset and the transition from MI to MII, when the first polar body is extruded (Fig. 2). CDK1 activity is then quickly reestablished and sustained during MII arrest [12,16–19]. Precise regulation of CDK1 activity during meiotic maturation involves both the modulation of cyclin B1 levels as well as phosphorylation of the CDK1 kinase domain at specific amino acid residues [20,21] Studies show that entry into metaphase is driven by cyclin B1 synthesis and

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CDK1 activation, whereas anaphase onset is correlated with ubiquitin-mediated cyclin B1 degradation. In fully grown oocytes, GV arrest is sustained in part due to constant degradation of cyclin B1 by the anaphase-promoting complex/cyclosome (APC/C)-Cdh1, which keeps CDK1 activity low [22]. Translational control for the availability of cyclin B1 also plays an important role [23,24]. In addition to regulating cyclin B levels, during prophase arrest CDK1 is maintained in an inactive state by phosphorylation on Thr14 and Tyr15 mediated by the inhibitory kinases MYT1 and WEE1 [25]. In mouse oocytes, Wee1B was identified as a key inhibitory kinase that phosphorylates CDK1 at Tyr15 [26]. Maintenance of this phosphorylated state is crucial for the prolonged prophase arrest in growing oocytes [27]. Just before the resumption of meiosis in fully grown oocytes, these sites on CDK1 are dephosphorylated by the dual specificity phosphatase, CDC25, leading to CDK1 activation [28]. Temporal changes in the subcellular distribution of cdc25 as well as the inhibitory kinases, Wee1b and Myt1, contribute to the regulation of meiotic resumption [29,30]. Notably, oocytes from mice with a null mutation for Cdc25b are ovulated at the GV stage and fail to

FIG. 2 (Top) Diagramatic representation of the major steps in the progression of nuclear maturation in mammalian oocytes. (Bottom) Diagramatic representation of the activity of maturation-promoting factor (MPF, solid line) and mitogen-activated protein kinase (MAPK, dashed line) during nuclear maturation. MPF activity is defined by the activation of the CDK1 protein kinase, which is regulated by de-phosphorylation on Thr14 and Tyr15 (green, active state). Decreased CDK1 activity is observed upon anaphase onset, owing to rephosphorylation (red, inactive state) of CDK1 as well as degradation of the Cyclin B (CYB) regulatory subunit. MPF activity is reestablished during MII arrest.

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resume meiosis [31]. Thus, establishing that Cdc25b is the key phosphatase essential for CDK1 activation at the G2/M transition in mouse oocytes. Appropriate mechanisms are essential to regulate CDK1 activity following the resumption of meiosis [32] and also maintain mature eggs arrested at MII until fertilization. The molecular basis for MII arrest was first investigated in Xenopus oocytes. A seminal experiment by Masui and Markert demonstrated that the transfer of cytoplasm from a mature MII egg into one blastomere of a two-cell embryo blocked cell division and maintained the injected blastomere in metaphase [12]. This cell cycle arresting activity, termed “cytostatic factor” (CSF), is necessary to sustain meiotic MII arrest in unfertilized vertebrate eggs. CSF activity is detected in oocytes at MII and disappears soon after fertilization [16,17,33]. Subsequent studies revealed that CSF functions primarily to stabilize CDK1 activity. Several signaling pathways play a vital role in maintaining MII arrest by sustaining sufficient Cyclin B levels in the oocyte, including Mos/ MAPK [34–36] as well as the anaphase promoting complex/cyclosome (APC/C) inhibitor Emi2 [37,38]. In addition to Cyclin B regulation, cdc25A [39] activity participates in MII arrest by promoting CDK1 activation.

Meiotic Competence The ability (competence) to resume and complete meiotic maturation is acquired during oocyte growth [5]. While fully grown oocytes can spontaneously resume meiosis in culture, smaller oocytes from preantral follicles cultured under the same conditions do not mature. Acquisition of competence to undergo GVB is correlated with the stage of follicle development as well as oocyte size. Oocytes first acquire the ability to undergo GVB when partially grown, but are not yet capable of completing maturation and prematurely arrest at MI. It is only with further development that oocytes can undergo the first meiotic division and progress to MII to become mature eggs [5,40]. The prolonged prophase arrest in growing oocytes is attributed, in part, to limited levels of key kinases, such as CDK1 as well as the maintenance of CDK1 in a phosphorylated (inactive) state [27]. In turn, the resumption and completion of meiosis in fully grown oocytes necessitates that key proteins essential for nuclear maturation are available and active. Levels of CDK1 and cyclin B1 proteins increase dramatically as oocytes acquire competence to undergo GVB [21,41,42]. However, total transcript levels are not significantly altered and point to the importance of translational control for protein concentrations at this stage [21,23,24]. Moreover, CDK1 and cyclin B1 redistribute and localize to the GV as oocytes gain the capacity to undergo GVB [29,43].

Importantly, deletion of Cdc25b, the phosphatase that activates CDK1, in mice leads to failure of GVB in fully grown oocytes [31]. These studies support that competence to resume meiosis is regulated not only by sufficient levels of key regulatory molecules, such as CDK1, but also their precise subcellular distribution and activation within the oocyte.

Maintenance of Prophase Arrest in Oocytes From Antral Follicles Fully grown oocytes with sufficient levels of regulatory molecules must control CDK1 activity to sustain prophase arrest until the LH surge. Identification of the specific molecules that promote meiotic arrest has been the subject of extensive investigation. Early oocyte culture and biochemical analyses were fundamental in delineating key signaling pathways that regulate meiotic arrest. In turn, genetic studies confirmed and further defined the underlying cellular and molecular mechanisms. Spontaneous maturation observed in oocytes upon release from the follicular environment [6,7] initially advanced the hypothesis that somatic cells in antral follicles help to maintain oocytes in prophase arrest. It was proposed that granulosa cell-derived signals might be transmitted to the oocyte via gap junctions [44,45]. In addition, factors present in ovarian follicular fluid were thought to play an important role [46]. Follicular fluid contains high concentrations of the purine, hypoxanthine, that can maintain oocytes at the GV stage [47–49]. Interestingly, hypoxanthine exhibits cyclic adenosine 30 50 -monophosphate (cAMP) phosphodiesterase inhibitory activity [50–52]. Since, phosphodiesterase (PDE) enzymes normally hydrolyze and lower cAMP levels, these studies underscored a key role for cAMP in sustaining meiotic arrest in oocytes. cAMP Produced in the Oocyte Promotes Prophase Arrest It is now well established that high levels of cAMP within the oocyte play a central role in preventing CDK1 activation and sustaining prophase arrest. Investigations demonstrated that membrane permeable analogs of cAMP promote meiotic arrest in vitro [53,54] and, that treatment of denuded oocytes with a direct agonist of adenylyl cyclase (AC), which catalyzes the formation of cAMP from ATP, delays the onset of GVB [55,56]. Moreover, general PDE inhibitors also sustain high levels of cAMP by blocking its breakdown and, hence, prevent GVB [57]. PDE3 has been identified as the major PDE expressed in oocytes from many species including human, and PDE3-specific inhibitors effectively prevent GVB in vitro [58–62]. Similarly, injection of rats with a PDE3-specific inhibitor results in the ovulation of oocytes arrested at prophase-I [42,63]. Moreover, PDE3A

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deficient female mice (Pde3a/) ovulate GV-arrested oocytes with high cAMP levels and are infertile [64]. These studies confirmed that sustained cAMP within the oocyte is crucial for meiotic arrest, and that PDE3 is a key regulator of cAMP levels and nuclear maturation. The steady-state level of cAMP relies on the balance of its synthesis and degradation, and it is important to understand how sufficient GVB-inhibiting levels of cAMP are sustained in oocytes. A central question initially focused on whether cAMP is generated within the oocyte itself or produced by the surrounding granulosa cells and then transported to the oocyte [44,51,65,66]. Key investigations have since determined that it is oocyte-derived cAMP, which is essential for the maintenance of meiotic arrest. Studies revealed that oocytes synthesize cAMP via stimulation of G-protein-coupled receptors (GPR) present in their plasma membrane. Adenylate cyclase (AC) genes and proteins are expressed within the oocyte, and mouse oocytes deficient in AC3 (Adcy3/) show defects in the maintenance of GV arrest [67]. Notably, studies demonstrated that the constitutively active G-protein coupled receptor 3 (GPR3) stimulates G protein (Gs) to activate adenylate cyclase, which in turn catalyzes formation of cAMP from ATP in oocytes [68–71]. Blocking Gs in follicle-enclosed oocytes prevents cAMP synthesis in the oocyte and promotes the resumption of meiosis [68,72]. Similarly, mouse oocytes lacking Gpr3 undergo GVB within the ovarian follicle [69,73]. This important mechanism is conserved and functional in human oocytes [74]. To sustain meiotic arrest cAMP activates a key downstream target, protein kinase A (PKA), which regulates CDK1 activity within the oocyte. Microinjection of the catalytic subunit of PKA into oocytes blocks spontaneous resumption of meiosis [75]. Moreover, inhibition of PKA signaling in Pde3A deficient oocytes promotes GVB [64], further supporting that prophase arrest by cAMP is mediated through PKA activation. It is important to note that PKA does not directly target CDK1. Instead PKA phosphorylates and activates the Wee1B kinase that normally phosphorylates and inactivates CDK1 [26,76]. In addition, PKA phosphorylates the phosphatase CDC25B [77]. Phosphorylated CDC25 is bound to 14-3-3 proteins and sequestered within oocyte cytoplasm, such that it cannot be dephosphorylated and activate CDK1 in the nucleus [30,78]. These studies established that cAMP signaling maintains CDK1 in an inactive state, through PKA-mediated inhibition of CDC25B and activation of WEE1B, to promote meiotic arrest in the oocyte. In tandem with cAMP/PKA-mediated inactivation, CDK1’s activity is further regulated by the availability of its regulatory subunit, cyclin B1. Continual degradation of cyclin B1 by the anaphase-promoting complex/cyclosome (APC/C) also promotes prophase arrest [22]. Disruption of this process leads to accumulation of cyclin B1 and resumption of meiosis [79].

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cGMP From Granulosa Cells Regulates cAMP Levels Within the Oocyte While the oocyte actively produces cAMP to maintain meiotic arrest, the surrounding granulosa cells play a pivotal role in regulating oocyte cAMP levels and meiotic resumption. Pioneering studies [6] showing that oocytes spontaneously resume meiosis when released from the follicle, first suggested that somatic cells likely provide an inhibitory signal to sustain oocyte arrest. Investigations have since addressed the potential mechanisms and identified an important link between cAMP and cyclic guanosine monophosphate (cGMP) signaling in the oocyte and granulosa cells in the follicle. Similar to cAMP, cGMP is reduced in the oocyte following removal from the follicle and resumption of meiosis as well as LH stimulation [57,80–82]. Cyclic GMP is produced in granulosa cells and functions as a potent inhibitor of PDE3 activity in the oocyte [82,83]. Importantly, disruption of gap junction communication between the oocyte and granulosa cells reduces both cGMP and cAMP in the oocyte, promoting the resumption of meiosis [84,85]. Lowering cGMP in the oocyte leads to the resumption of meiosis, but not when a PDE3 inhibitor is present. These studies confirm that granulosa cells support high cAMP in the oocyte by producing and transferring cGMP to the oocyte via gap junctions. In turn, cGMP inhibits PDE3 activity within the oocyte, blocking cAMP hydrolysis and thereby sustaining high cAMP levels for meiotic arrest [64,82,85]. Considering the function of cGMP, control of its availability in granulosa cells is essential for maintaining meiotic arrest. Studies identified a central role for the natriuretic peptide precursor C (NPCC) and its receptor, natriuretic peptide receptor 2 (NPR2), in regulating cGMP synthesis in granulosa cells. In mice, mural granulosa cells express Npcc while the cumulus cells surrounding the oocyte express its receptor, Npr2, a guanylyl cyclase. The NPCC peptide inhibits spontaneous GVB in cumulus enclosed, but not in denuded, oocytes in culture. Notably, oocytes from Npr2 or Npcc-null mice resume meiosis spontaneously in vivo [86] and cGMP levels are undetectable in antral follicles from female mice with a loss of function mutation in Npr2 [87]. These studies established that NPPC produced by mural granulosa cells activates NPR2 receptor in cumulus cells to promote cGMP synthesis. Interestingly, oocyte-derived paracrine factors promote cumulus cell expression of Npr2 mRNA and regulation of cGMP production, underscoring an important bidirectional communication loop [86,88]. Thus, the oocyte supports meiotic arrest not only by producing cAMP, but also by promoting NPR2 receptor expression in cumulus cells to generate cGMP to inhibit oocyte PDE3 activity.

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The Induction of Nuclear Maturation by LH LH is the physiological signal that promotes ovulation and initiates the resumption of meiosis in oocytes. Specific LH receptors are expressed in mural granulosa cells, with little or no expression detected in cumulus cells or the oocyte [89,90]. This poses the question as to how LH signaling at the outer mural granulosa cells promotes meiotic resumption in the oocyte. Early studies propose that LH-mediated disruption of gap junction communication between the oocyte and surrounding granulosa cells [44,91] plays a significant role in instigating oocyte nuclear maturation. Incubation of follicle-enclosed oocytes with a gap junction disruptor (carbenoxolone) lowers oocyte cAMP levels and leads to meiotic resumption [84], supporting the importance of gap junction communication. Notably, discovery that cGMP produced in granulosa cells is transferred, via gap junctions, to the oocyte where it blocks PDE3 activity to sustain high cAMP levels provided critical insight into the nature of signaling between granulosa cells and the oocyte to regulate meiotic arrest [64,82,85]. In response to LH, cGMP levels decrease significantly in granulosa cells as well as the oocyte. Elegant studies revealed that a rapid LH-mediated decrease in cGMP is first observed in the mural granulosa cells, which express LH receptors. Lower cGMP is subsequently observed in the cumulus cells and then the oocyte [92] as less cGMP is available and transferred from the granulosa cells. The decrease in cGMP levels mediated by LH is attributed to lower cGMP synthesis by NPR2 [93–95]. Additionally, increased breakdown of cGMP by the phosphodiesterase PDE9 [96] also contributes to LH-mediated attenuation of cGMP within the follicle. Collectively these studies demonstrate that LH signaling promotes a rapid decline in mural granulosa cell cGMP levels, limiting its availability for transfer to the oocyte. In turn, low cGMP in the oocyte precludes inhibition of PDE3 activity, thereby instigating the hydrolysis and reduction of cAMP levels in the oocyte. The significant decrease in cAMP leads to the activation of CDK1, initiating GVB. Hence, the maintenance of prophase arrest as well as meiotic resumption is contingent on crucial bidirectional and coordinated communication between the oocyte and somatic granulosa cells within the follicle.

EPIGENETIC MATURATION In mammalian oocytes, the process of epigenetic maturation takes place during a critical window of postnatal oocyte growth and differentiation of the ovarian follicle. Human oocyte growth from the primordial stage to the fully grown preovulatory stage requires 3–4 months, while mouse oocytes require only 3 weeks to complete

this process and reach the preovulatory stage in preparation for meiotic maturation [97]. Shortly after birth, and coincident with activation of primordial follicles, ovarian granulosa cells establish an intimate bidirectional communication with the growing oocyte that is essential for the synthesis and storage of maternal mRNA, accumulation of cell cycle regulatory proteins, cytoplasmic organelles, and formation of the zona pellucida [97,98]. Granulosa cells modulate the process of transcription and chromatin remodeling in the oocyte genome [99,100], play an active role in the transport of RNA molecules to the oocyte through the establishment of intercellular communication [101], and can regulate translation of maternal mRNA stores [102]. In turn, functional differentiation of chromatin structure in the oocyte is essential for the control of gene expression, establishment of maternal genomic imprinting, maintenance of genome stability as well as the acquisition of both meiotic and developmental competence [103,104]. Chromatin structure and function during oocyte growth is regulated by multiple and hierarchical epigenetic modifications established during developmental transitions or in response to endocrine or environmental stimuli [104]. Epigenetic modifications were initially defined as stable and heritable chromatin modifications that influence gene expression without inducing any changes in the underlying DNA sequence. However, a more contemporary definition encompassing the dynamic nature of myriad chromatin marks and their interactions in the mammalian genome considers epigenetics as “the structural adaptation of chromosomal regions so as to register, signal or perpetuate altered activity states.” Thus, the initial definition has now been expanded to also take into consideration a number of transient modifications that are essential for DNA repair or that may be unique to a specific cell cycle stage in addition to stable changes inherited through mitosis or meiosis [105]. Examples of epigenetic modifications in the mammalian genome include DNA methylation, histone posttranslational modifications, such as histone acetylation, histone methylation, histone phosphorylation and poly (ADP) ribosylation, the incorporation of histone variants, such as histone H3.3 or the centromere specific histone H3 variant CENP-A. Epigenetic modifications are highly dynamic and may work in synergy or antagonistically in order to trigger the changes in gene expression that are required in response to a wide range of differentiation and/or environmental stimuli. Importantly, they are capable of acting at the single gene level, within the context of a specific genomic or chromosomal domain, such as centromeres or telomeres or genome-wide at the level of large-scale chromatin remodeling [106]. The range of biological mechanisms under epigenetic control is diverse, and involves some of the most fundamental principles of genome organization. Epigenetic

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modifications regulate genome-wide topological organization and thus have a direct effect on gene expression and cell fate. Importantly, due to their ability to act at many different levels of nuclear organization, epigenetic changes may regulate genome architecture from chromatin fiber compaction to global effects on the threedimensional organization of the nucleus [106]. The evidence obtained until now indicates that in the oocyte genome, epigenetic modifications are required for the control of transcription, establishment of maternal DNA methylation patterns, nuclear architecture, and maintenance of chromosome stability [104,107–111]. Notably, many epigenetic modifications exhibit a germ cell-specific function in order to remodel the genome generation after generation as well as to fulfill the unique requirements of gamete formation and accurate chromosome segregation during meiosis [104,112–115].

Establishment of Epigenetic Modifications During Oogenesis DNA methylation is one of the most widely studied epigenetic modifications in the mammalian genome. Addition of a methyl group [CH3] at the carbon 5 position of specific cytosine DNA bases is catalyzed by DNA methyl transferase proteins (DNMT-1, DNMT3A, and DNMT3B) resulting in the formation of 5-methyl cytosine. In mouse germ cells, the interaction of DNMT3A with its catalytic activity DNMT3L creates a gamete-specific DNA methylation mark with profound consequences for the control of gene expression, normal embryonic development, placental differentiation and fetal growth [116]. In most mammalian cell types DNA methylation occurs mainly at cytosine-guanine dinucleotides called CpG islands that frequently colocalize with gene promoters and regulatory regions [116,117]. Notably, in mammalian oocytes DNA methylation can also occur at non-CpG islands and in fact show a clear enrichment within gene bodies [118–120]. Non-CpG methylation is abundant in embryonic stem cells, germ cells, and the brain and may constitute a new layer of genome regulation although its function remains to be determined [119,120]. Evidence obtained from DNMT3A/ DNMT3L knockout mice indicates that oocytes can mature, progress through meiosis, and become fertilized in the absence of DNA methylation at maternally imprinted CpG islands, also called germ line differentially methylated regions (gDMRs). However, the patterns of DNA methylation established during oogenesis play a critical role in the regulation of gene expression during subsequent embryo development as embryos from DNMT3A/DNMT3L-deficient oocytes die by E10.5 due to absence of maternal methylation imprints, abnormal allelic transcription, and impaired trophoblast

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development [121]. Importantly, a growing body of evidence indicates that DNA methylation during oogenesis may also have a profound effect in regulating transgenerational epigenetic inheritance [121]. The ontogeny and mechanisms that establish DNA methylation patterns during oogenesis have been the subject of intense investigation [119,122,123]. Elegant nuclear transfer experiments in combination with DNA methylation analysis suggested that the establishment of de novo methylation patterns during oogenesis begins when the oocyte reaches 50 μM in diameter in preantral follicles around day 16 of postnatal development in the mouse [122–124]. Establishment of de novo methylation occurs on a locus-by-locus basis during subsequent stages of oocyte growth and is completed before global transcriptional repression takes place in fully grown, preovulatory oocytes within graafian follicles on day 21 of postnatal development in female mice [123]. Initial bisulfite DNA methylation assays revealed that establishment of maternal DNA methylation marks takes place asynchronously at different imprinted genes [122,124,125]. More recently, powerful strategies such as genome-wide DNA methylation and single nucleotide sequencing revealed the onset of de novo DNA methylation as early as day 10 of postnatal development in mouse oocytes [123] and have begun to shed important mechanistic insight into this critical process. Current evidence indicates that both transcription and histone modifications across gDMRs regulate the placing and time of establishing de novo methylation marks at defined loci [121,126,127]. Importantly, the effect of transcription on the establishment of DNA methylation seems to be unique to the oocyte [121]. However, changes in the rates of transcription are not sufficient to establish de novo methylation of imprinted genes. Instead chromatin remodeling at CpG islands is crucial to regulate the timing and progressive establishment of DNA methylation marks at different imprinted gDMRs during oogenesis, and may directly link the process of oocyte growth with the progressive establishment of DNA methylation at specific gDMRs [128]. Transcription may facilitate a chromatin configuration that facilitates the recruitment [128] of DNA methyl transferases (DNMT3A/DNMT3L) to establish maternal methylation marks [126]. Several lines of evidence indicate that functional interactions between DNMT3A/DNMT3L regulate the establishment of DNA methylation at gDMRs in mouse oocytes [129–131]. However, this mechanism requires the action of the histone demethylase KDM1B to remove a histone methylation mark (H3K4me2), which interferes with de novo methylation. Removal of this mark is indispensable for the establishment of DNA methylation at many, albeit not all, gDMRs suggesting that alternative pathways may also exist for different gDMRs [132]. The mechanisms of KDM1B function remain to be determined. However, it

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has been suggested that KDM1B may induce demethylation of H3K4me2 and potentially open the chromatin configuration to recruit the DNA methylation machinery to selected loci [132]. Chromatin immunoprecipitation (ChIP-seq) studies confirmed that the presence of H3K4me2 interferes with DNA methylation and must first be removed by KDM1B to establish DNA methylation at oocyte gDMRs, while permissive H3K36me3 marks increase in growing oocytes as de novo methylation is established [110]. This model implies that different histone modifications are likely required at different genomic locations [121,126,127]. The precise mechanisms by which transcription regulates chromatin remodeling remain to be determined as well as the functional interactions of several chromatin-binding proteins that are critical for the maintenance of DNA methylation patterns in oocytes and embryos, such as the maintenance DNA methylase UHRF1 [133], the KRAB zinc finger protein ZFP57 [134], Sall4 [135] also a zinc finger protein, the transcription factor TRIM28 [136], and the histone H3.3 chaperone HIRA [109]. Perhaps one of the most striking observations from genome-wide sequencing studies conducted by several independent laboratories is the unique nature of the oocyte methylome, consistent with the presence of germ-cell specific mechanisms for epigenetic regulation [108–111]. Indeed, oocyte transcriptome and DNA methylome analyses indicate that in contrast with most somatic cells analyzed until now, almost two-thirds of all methylcytosines occur in a non-CpG context and are clearly enriched at gene bodies. This reveals that the oocyte genome is compartmentalized into large-scale hypermethylated and hypomethylated domains in which transcription regulates the establishment of almost 90% of the oocyte methylome. Thus, DNA methylation is predominant at expressed gene bodies in fully grown oocytes [119,120,127]. Importantly, whereas promoter DNA methylation has been negatively associated with gene expression in many cell types, studies indicate that gene body methylation is positively correlated with gene expression in the mammalian oocyte [119,120,127]. Another remarkable epigenetic feature in mammalian oocytes is the unusual genomic distribution of H3K4me3. Although generally considered as a transcriptionally permissive histone modification, genome-wide chromatin immunoprecipitation studies (ChIP-seq) indicate that the relationship of H3K4me3 with transcription is far more complex than initially thought. In fact, the mouse oocyte has proven an invaluable model to uncover the presence of both transcription-dependent and transcription-independent mechanisms for H3K4me3 deposition [137]. For example, in embryonic stem cells and somatic cells H3K4me3 is preferentially enriched at transcription start sites (TSS) of active genes where it results in a narrow high peak of H3K4me3 following

ChIP-seq analysis. In contrast, during the oocyte to embryo transition the H3K4me3 mark is present in large genomic regions spanning more than ten thousand base pairs, with low signal density and found distant from the TSS. This unique pattern has been termed broad, or noncanonical, H3K4me3 to emphasize its difference with the clearly delineated peaks of H3K4me3 associated with transcription start sites in somatic and ES cells [138–140]. The mechanisms regulating broad H3K4me3 patterns and its effect on transcription or DNA methylation at specific compartments in the oocyte genome are not fully understood. However, use of a transgenic mouse model with an oocyte-conditional deletion of the histone methyltransferase (MLL2), known to regulate H3K4me3 [141] in combination with ultra-low input ChIP-seq analysis confirmed the presence of broad H3K4me3 patterns in fully grown oocytes [137]. Interestingly, in the nongrowing oocyte on day 5 of postnatal development H3K4me3 is restricted to transcribed gene promoters and is dependent on transcription. With further oocyte growth, accumulation of H3K4me3 becomes independent of transcription and it spreads throughout thousands of bivalent domains and broad distal domains. This noncanonical distribution is regulated by MLL2 function. Thus, there are two independent mechanisms of H3K4me3 deposition in the oocyte genome, with MLL2 specifically targeting unmethylated CpG-rich regions in a transcription independent manner. Notably, loss of MLL2 had no effect on transcription-coupled H3K4me3 deposition at TSS, DNA methylation or the oocyte transcriptome [137]. The zinc finger protein SALL4 has also been recently involved in the regulation of H3K4me3 patterns during oocyte growth as loss of SALL4 induced and overexpression of the histone de-methylase KDM5B with a subsequent reduction of H3K4me3 levels [135]. So what are the biological implications of broad H3K4me3 deposition patterns in the oocyte genome? Two intriguing observations provide some initial insight into the potential function of broad H3K4me3 genome deposition although this remains to be directly demonstrated. First, overexpression of the histone demethylase KDM5B reduced broad H3K4me3 distribution and reactivated transcription in preovulatory oocytes [138], which are known to undergo global transcriptional silencing in preparation for meiotic resumption [104]. Loss of MLL2 also prevented programmed transcriptional silencing in the oocyte suggesting that MLL2-dependent deposition of broad H3K4me3 domains may be required for global transcriptional silencing [137,141]. Second, removal of broad H3K4me3 domains is required for normal zygotic genome activation and suggests that this may be an important epigenetic mechanism for regulation of global transcriptional activity during the oocyte to embryo transition [138–140].

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Histone modifications have a direct impact in the regulation of chromatin structure and function [142]. By exerting changes in nucleosomal organization or through large-scale chromatin remodeling at the chromosomal level, histone posttranslational modifications have the potential to regulate the degree of chromatin condensation and establish either a transcriptionally permissive or a transcriptionally repressive chromatin environment [143,144]. The levels of both methylation and acetylation at different lysine residues of histone H3 and histone H4 increase during oocyte growth, concomitantly with the levels of expression of several histone acetyl transferases and histone methyl transferases [145]. However, we have only a very limited mechanistic understanding concerning the functional interaction of most of these modifications, and an increasingly complex “meiotic histone code,” with the control of transcriptional activity and large-scale chromatin remodeling in the oocyte genome [104,115,145]. Recent studies indicate that chromatin structure during oogenesis is highly dynamic and underscore the importance of continuous deposition and replacement of histone H3.3 and H4 by the histone chaperone HIRA, which is essential for chromatin homeostasis, the control of gene expression, the establishment of de novo methylation patterns in the oocyte and ultimately, female fertility [109]. Depletion of HIRA in primordial oocytes causes extensive oocyte death due to lack of H3/H4 deposition, increased DNA accessibility and accumulation of DNA damage. The resulting chromatin displayed altered structure, increased DNAseI sensitivity as well as double strand DNA breaks [109]. Notably, a systematic analysis of genome-wide histone modifications using a sensitive chromatin immunoprecipitation and DNA sequencing (ChIP-Seq) strategy revealed that histone modifications are established before the onset of de novo methylation and have a direct impact in the subsequent acquisition of oocyte-specific DNA methylation patterns [110]. Thus, histone modifications shape the oocyte methylome by directing the methylation machinery to specific genomic regions. Histone methylation at lysine 36 (H3K36me3) can recruit DNMT3A in vitro, suggesting that a similar mechanism may function in vivo [121]. However, different histone modifiers are likely required at different genomic locations. For example, as mentioned above, some CpG islands with high H3K4me2 and H3K4me3 may require active removal of this mark, while at different GpG islands deposition of H3K36me3 may be sufficient to establish DNA methylation [110,121]. Notably, a new type of noncanonical imprinting mechanism mediated by the inheritance of oocyte-specific H3K27me3 has been recently described in mouse oocytes that is critical for the regulation of a subset of genes whose imprinting is regulated through a DNA methylation-independent mechanism [146]. Other histone modifications are essential to regulate entire

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chromosomal domains. For example, histone trimethylation at lysine 9 (H3K9me3) correlates with DNA methylation at heterochromatin domains. Prominent histone modifications, such as H3K9me3, also play a critical role in regulating the function of large-scale chromosomal domains, such as centromeres and telomeres and together with H4K20me3 are essential for the formation of constitutive heterochromatin the regulation of largescale chromatin remodeling and maintenance of chromosome stability [104].

Large-Scale Chromatin Remodeling and Global Transcriptional Silencing in the Oocyte Genome The nucleus (GV) of most mammalian oocytes is subject to a striking process of large-scale chromatin remodeling in preparation for the resumption of meiosis. This process occurs simultaneously with global silencing of transcriptional activity and is strictly required for the acquisition of both meiotic and developmental potential as well as for the maintenance of chromosome stability during the oocyte to embryo transition [104]. Notably, in order to cope with the unique requirements of chromosome dynamics during meiosis, specialized mechanisms are set in place for the control of large-scale chromatin structure in the female gamete [104,112,114,115]. Large-scale chromatin remodeling is defined as a series of genome-wide changes in nuclear architecture that can be recognized at the level of specific chromosomes or chromosome domains such as centromeres [147]. Growing oocytes within late preantral follicles on day 17 of postnatal development in the mouse exhibit a large nucleus characterized by highly decondensed chromatin and several prominent heterochromatin domains or chromocenters that can be found dispersed throughout the entire GV, a configuration known as the “nonsurrounded nucleolus” or (NSN) [148–150]. These oocytes exhibit high levels of global transcriptional activity as sensitive transcription run-on assays can detect abundant nascent transcripts in the nucleoplasm within 20 min following RNA labeling [99]. In addition, western blot analysis revealed high levels of the active or hyperphosphorylated form of the RNA polymerase II enzyme [149]. With further oocyte growth and differentiation and coincident with the formation of early antral follicles that contain a fully grown preovulatory oocyte, the GV undergoes a striking, developmentally programed, large-scale chromatin remodeling event that is essential to confer the oocyte with the ability to complete meiotic maturation and pre-implantation development [99,149,151]. At this stage, changes in global or large-scale chromatin structure lead to a progressive condensation of DNA and the formation of a prominent perinucleolar heterochromatin rim known as the “surrounded nucleolus”

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(SN) configuration or karyosphere. This configuration is present in both human and mouse oocytes [148,149,152] as well as oocytes from several domestic species [104]. Notably, transcription run-on assays demonstrate that formation of the SN configuration in mouse and human oocytes occurs concomitantly with global repression of transcriptional activity most likely due to displacement of RNA polymerase II from the chromatin template before GV breakdown in preparation for meiotic resumption [99,149,151,153,154]. Previous studies indicate that oocytes that exhibit the NSN configuration are able to undergo GV breakdown and meiotic resumption but show an arrest at the metaphase I stage. In contrast, oocytes with the SN configuration have acquired the capacity to complete meiotic resumption, reach the metaphase II stage, and exhibit a higher developmental potential to the blastocyst stage [151,155]. Thus, formation of the SN configuration or karyosphere is critical for the timely progression of meiotic maturation. Indeed, several lines of evidence indicate that large-scale chromatin remodeling in the oocyte genome is essential for the acquisition of meiotic and developmental competence [103,104,107,156]. However, we have only a limited mechanistic understanding of the potential developmental and signaling mechanisms regulating large-scale chromatin remodeling in the mammalian oocyte genome. The initial evidence suggesting that programmed chromatin remodeling in the oocyte genome may be regulated, at least in part, by signaling pathways emanating from companion granulosa cells was obtained using a unique experimental system designed for the culture of mouse oocyte-granulosa cell complexes obtained from preantral follicles [99]. Abnormal chromatin remodeling following culture of denuded oocytes that lack any functional interactions with granulosa cells suggested that developmentally regulated signal(s), presumably of paracrine origin from granulosa cells modulate chromatin remodeling and global transcriptional activity in mouse preovulatory oocytes [99]. Additional evidence obtained by independent laboratories support this model [100]. Together, the emerging picture suggests that communication between the oocyte and its companion granulosa cells mediated by the establishment of patent gap junctions is required for programmed chromatin remodeling as growing oocytes reach the preovulatory stage. However, the cell signaling pathways under granulosa cell control remain to be determined [104]. Notably, although large-scale chromatin remodeling into the SN configuration and global transcriptional silencing in preovulatory oocytes are both essential for the acquisition of developmental potential, these two mechanisms can be experimentally dissociated suggesting that they are regulated through different pathways [103]. Two lines of evidence revealed the existence of independent pathways. Both pharmacological manipulation of chromatin

structure as well as analysis of transgenic mouse models indicate that although temporally linked in wild type preovulatory oocytes, chromatin remodeling into the SN configuration and global transcriptional repression are independently regulated. First, treatment of wild-type preovulatory oocytes that exhibit the SN configuration with the histone deacetylase inhibitor trichostatin A (TSA) induced the formation of highly decondensed chromatin and disrupted the maintenance of the karyosphere or SN configuration, yet global transcriptional activity remained silenced. Second, analysis of transgenic oocytes with a targeted deletion of the nucleolar protein nucleoplasmin 2 (Npm2) revealed that these oocytes fail to remodel chromatin into the SN configuration, yet global transcriptional repression as detected by transcription run-on assays occurs in a timely manner [103]. More recently, evidence has also been obtained for the presence of transgenic preovulatory oocytes that exhibit the SN configuration in which global transcriptional activity remains unabated. Two genetic mouse models have been particularly important to illustrate this process [141,157]. Targeted deletion of the histone H3K4 methyltransferase Mll2 resulted in the loss of trimethylation of Histone H3 at lysine 4 (H3K4me3) in preovulatory oocytes and lack of global transcriptional repression in spite of the presence of an SN configuration [141]. Moreover, targeted deletion of the dual bromodomain-containing protein BRWD1 prevented global transcriptional repression in >70% of preovulatory oocytes that exhibited a SN configuration [157]. These studies provide conclusive evidence for the existence of independent mechanisms for the control of chromatin remodeling and global transcriptional silencing in the mouse oocyte. In drosophila germ cells, the karyosome is an oocytespecific chromatin configuration, analogous to the karyosphere that is present in mammalian oocytes. Depletion of the nucleosomal histone kinase-1 (NHK-1) also known as VRK1 in Drosophila eggs causes striking epigenetic changes, such as loss of histone H2A phosphorylation (H2AT119ph), decreased histone H3 (H3K14ac), and histone H4 (H4K5ac) acetylation. Notably, loss of VRK1 in drosophila oocytes leads to abnormal chromatin remodeling and abnormal karyosome formation [158,159]. However, analysis of vaccinia related kinase (VRK1) knockout mouse oocytes showed only a mild chromosome segregation defect. Instead the most prominent phenotype observed in these oocytes was a complete absence of pronuclear formation and developmental arrest following in vitro fertilization [160]. Thus, further studies are required to unravel the signaling pathways involved in the developmental regulation of karyosphere formation in the mammalian oocyte genome. Formation of the karyosphere is a critical developmental transition at the culmination of oocyte growth. Notably, the specialized nuclear architecture acquired during

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the transition into the SN configuration is essential for the control of centromeric heterochromatin domains and the epigenetic control of centromere function [103]. Thus, large-scale chromatin remodeling in preovulatory oocytes is essential for both accurate chromosome segregation during meiotic maturation and for the maintenance of chromosome stability during the oocyte to embryo transition [103]. In somatic cells, large-scale chromatin remodeling is mediated by binding of ATPdependent chromatin remodeling proteins, histone variants such as CENP-A, as well as histone posttranslational modifications, such as histone acetylation or histone methylation at different lysine residues of the histone tail [142,147]. The interaction of multiple histone modifications provides structural and functional identity to individual chromosome domains such as centromeres and, therefore, has a major impact in the control of accurate chromosome segregation and chromosome stability. For example, centromeric heterochromatin formation is critically important to regulate centromere-microtubule interactions, homologous chromosome segregation during metaphase-I and sister chromatid separation during metaphase II [103,161]. The mechanisms regulating the “meiotic histone code” and its impact in large-scale chromatin structure, chromosome stability as well as maternal inheritance of epigenetic states to the early embryo are only beginning to be unraveled [115]. The evidence obtained until now indicates that some histone modifications required for the formation of transcriptionally repressive chromatin at centromeres, such as H3K9me3, are highly stable and remain associated with centromeric heterochromatin throughout meiosis [104]. In contrast, global histone acetylation exhibits highly dynamic patterns during meiotic resumption and prior to chromosome segregation [103,156,162,163]. For example, mouse oocytes exhibit high levels of histone H3 and H4 at most of the lysine residues of the histone tails resulting in wide-spread nuclear localization of these marks at the GV of fully grown, preovulatory oocytes. However, as the female gamete prepares to resume meiosis and shortly after GV breakdown the entire oocyte genome is subject to a wave of global histone deacetylation that removes most chromosome bound acetylation marks, such as H4K12ac and H4K5ac [103,156,163]. Global histone deacetylation is critical to regulate proper chromosome condensation, coordinate chromosome-microtubule interactions, and ensure accurate chromosome segregation [103,156,163]. Interfering with this process by incubating mouse oocytes with the histone deacetylase inhibitor trichostatin-A (TSA) affects the epigenetic control of centromere function by preventing the binding of vital centromeric proteins, such as the chromatin remodeling factor ATRX, and inducing the formation of highly elongated chromatids that frequently form improper attachment with microtubules

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resulting in severe chromosome segregation defects [103,156]. Exposure of metaphase II oocytes to TSA also leads to oocyte aneuploidy and subfertility in vivo [164]. In addition, the localization of the chromosome passenger complex to the centromere also requires global histone deacetylation [165]. The mechanisms responsible for regulating this important developmental transition are not fully understood. Notably, current evidence indicates that global histone deacetylation exhibits distinct strategies in female germ cells compared with somatic cells [103,163]. For example, in somatic cells histone deacetylase 3 (HDAC3) is the main enzyme regulating de-acetylation of H3 and H4 during mitotic chromosome condensation [166]. In contrast, histone deacetylase 2 (HDAC2) seems to be the critical deacetylase required for global deacetylation in mouse oocytes, specifically of histone H4 acetylation at lysine 16 (H4K16ac). In addition, the retinoblastoma-binding protein (RBBP7) has been recently involved in global histone deacetylation of H3K4ac, H4K8ac, H4K12ac, and H4K16ac but not H3K9ac or H3K14ac [165]. This suggests that different acetylation marks are removed from chromosomes by yet to be identified specialized histone deacetylase enzymes [115]. At the clinical level, reproductive aging is now widely recognized as a major risk factor for human oocyte aneuploidy [167]. Mouse oocytes retrieved from females of advanced reproductive age exhibit high levels of histone acetylation (H4K8ac and H4K12ac) at the metaphase II stage [164,168]. Importantly, a similar correlation between advanced maternal age, defective histone deacetylation and chromosome segregation defects has been reported in human oocytes [169]. In both human and mouse oocytes the “maternal age effect” remains as a virtual black box in the field of reproductive biology. However, this is perhaps one of the most critical issues affecting female fertility as the probability of ovulating a chromosomally abnormal egg increases up to 35% or more in females of advanced reproductive age [167,170]. Meiotic spindle defects are also an important component of mammalian oocyte aneuploidy during reproductive senescence [167,171] and it will be important to determine the specific mechanisms leading to improper chromosome microtubule interactions during meiosis. Microarray analyses to compare the transcriptome of young and old mouse oocytes have been instrumental in identification of key cellular pathways that are disrupted during oocyte aging. The most prominent pathways identified until now involve mechanisms regulating the spindle assembly checkpoint, kinetochore function, a deterioration of cohesion complexes as well as chromatin remodeling processes required for meiosis [172–176]. Collectively all these pathways contribute to a different extent to age-related subfertility and impaired embryonic development [177,178]. Importantly, the epigenetic maturation of centromeric heterochromatin

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domains may be particularly susceptible to both endocrine and environmental disturbances that result in wide-spread chromosome instability during the oocyte to embryo transition [157,179,180].

CONCLUSION Oocyte maturation stands uniquely at the nexus of developmental and reproductive biology. The processes encompassing oocyte maturation are essential for the transition from a gamete to an embryo competent to give rise to a healthy new individual. Moreover, these same processes represent the final stage in the generation of a highly differentiated cell, the egg. Excellent strides have been made to determine the molecular basis of oocyte maturation, yet many key questions remain to be resolved. An increased understanding of oocyte nuclear and epigenetic maturation is critical for the development of novel approaches to fertility control and reducing the incidence of birth defects, as well as facilitating efficient domestic animal production and preservation of endangered species.

Acknowledgments We thank Katelyn Snell and Brad Gilleland (Educational Resource Center, University of Georgia) for preparation of the artwork in Fig. 2. Funding support provided by NSF through the Research Center for Cell Manufacturing CMaT, The Regenerative Engineering and Medicine Center and an USDA-NIFA Animal Health Capacity Pilot Grant to Dr. De La Fuente. Funding support was also provided by NIH (HD086528 and HD0713330) to Dr. Viveiros.

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[166] Li Y, et al. A novel histone deacetylase pathway regulates mitosis by modulating Aurora B kinase activity. Genes Dev 2006; 20:2566–79. [167] Hassold T, Hunt P. To err (meiotically) is human: the genesis of human aneuploidy. Nat Rev Genet 2001;2:280–91. [168] Suo L, et al. Changes in acetylation on lysine 12 of histone H4 (acH4K12) of murine oocytes during maternal aging may affect fertilization and subsequent embryo development. Fertil Steril 2010;93:945–51. [169] van den Berg IM, et al. Defective deacetylation of histone 4 K12 in human oocytes is associated with advanced maternal age and chromosome misalignment. Hum Reprod 2011;(5):1181–90. [170] Hunt PA, Hassold TJ. Human female meiosis: what makes a good egg go bad? Trends Genet 2008;24:86–93. [171] Vialard F, et al. Evidence of a high proportion of premature unbalanced separation of sister chromatids in the first polar bodies of women of advanced age. Hum Reprod 2006;21:1172–8. [172] Chiang T, Duncan FE, Schindler K, Schultz RM, Lampson MA. Evidence that weakened centromere cohesion is a leading cause of age-related aneuploidy in oocytes. Curr Biol 2010;20: 1522–8. [173] Hodges CA, Revenkova E, Jessberger R, Hassold TJ, Hunt PA. SMC1beta-deficient female mice provide evidence that

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12 Oocyte Meiotic Maturation Heng-Yu Fan, Qing-Yuan Sun INTRODUCTION In mammals, oocytes begin to arrest at prophase of meiosis I (prophase I) at embryonic or perinatal stages. Oocytes arrested at prophase I contain a large nucleus covered by a nuclear envelope which is also known as the germinal vesicle (GV). The lengthy, complex, and discontinuous meiotic process that gives rise to a mature egg is called oocyte meiotic maturation. This process encompasses four developmental programs that are essential for the production of an egg competent to undergo fertilization and embryogenesis: (1) Nuclear maturation. Resumption of meiosis occurs in response to the surge of luteinizing hormone (LH) or upon the mechanical release of the oocyte from the ovarian follicle and subsequent in vitro culture in a suitable medium. Germinal vesicle breakdown (GVBD) is the first morphological sign of meiotic resumption. Following GVBD, meiosis I spindles are formed and bivalent chromosomes align along their chiasmata. Chiasmata are resolved once all bivalent chromosomes aligned well on middle plate and come under microtubule tension leading to anaphase I. Meiosis I is completed by emitting a polar body (PB) containing one set of chromosomes. The oocytes subsequently enter into the second round of meiosis but arrest again at metaphase of meiosis II (MII) until fertilization. (2) Cytoplasmic molecular maturation. Oocyte developmental competence is the ability of the mature oocyte to be fertilized and subsequently drive early embryo development. Developmental competence is acquired by completion of an oocyte maturation process that includes cytoplasmic molecular changes. Given that maturing oocytes are transcriptionally quiescent, they depend on posttranscriptional regulation of stored transcripts for protein synthesis, which is largely mediated by translational repression and deadenylation of

The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00012-1

transcripts within the cytoplasm, followed by recruitment of specific transcripts in a spatiotemporal manner for translation during oocyte maturation. (3) Cytoplasmic organelle maturation. Ultrastructural analysis and immunofluorescent staining have shown that mitochondria, ribosomes, endoplasmic reticulum (ER), cortical granules, and the Golgi complex assume different positions during the transition from the GV stage to metaphase II stage and their improper distribution and function affect oocyte cytoplasmic maturation quality. (4) Epigenetic maturation. This process involves de novo DNA methylation that silences maternal imprinted genes, exchanges of histone variants, and histone modification that support genome remodeling and gene transcription, during oogenesis and preimplantation development. Studies on mammalian systems have suggested the existence of intergenerational (between) or transgenerational (across multiple) epigenetic inheritance of acquired traits [1]. Proper epigenetic modifications during oocyte maturation are required for gene regulation or other chromatin-based processes in the next generation.

OOCYTE NUCLEAR MATURATION Mechanism of Oocyte Meiosis Arrest at Prophase I Mammalian oocytes within fully grown antral follicles remain arrested at prophase I and do not resume meiosis until there is a preovulatory surge of LH. Nonetheless, fully grown oocytes also spontaneously resumed meiosis when freed from the follicle and cultured in vitro (Fig. 1A). This observation led to the hypothesis that the follicular microenvironment prevents the resumption of oocyte meiosis and that the LH surge removes this

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FIG. 1 Schematic representation of the mammalian oocyte meiotic maturation process. (A) Within the follicle, oocytes remain arrested at prophase I and are characterized by a germinal vesicle (GV) in which the chromosomes remain decondensed. A preovulatory luteinizing hormone (LH) surge under physiological conditions or mechanical release of the oocyte from follicle followed by in vitro culture causes germinal vesicle breakdown (GVBD); chromosomes start to condense and spindle fibers appear. Subsequently, chromosomes condense and align at the metaphasic plate of the bipolar spindle when the oocyte reaches metaphase I. Meiosis I is completed by extruding a polar body containing one set of chromosomes. The second set of chromosomes is retained in the oocyte, which is now referred to as a secondary oocyte (or an egg). The mature egg remains arrested at metaphase II until fertilization. (B) Diagram of the activity of maturation promoting factor (MPF) (solid line) and mitogen-activated protein kinase (MAPK; dashed line) during nuclear maturation. MPF activity is defined by the activation of the cyclin-dependent kinase-1 (CDK1) protein kinase component of MPF, whose activity is regulated by dephosphorylation at Thr-14 and Tyr-15 (green). Loss of CDK1 activity is brought about at anaphase I or II by the rephosphorylation (red) of CDK1 and the proteolytic degradation of the cyclin B (CYB) regulatory component of MPF.

inhibitory factor rather than providing any positive stimulus for the resumption of meiosis. Reentry of meiosis depends on the activity of maturation promoting factor (MPF) in oocyte. MPF is a complex of a catalytic cyclin-dependent kinase-1 (CDK1) subunit and its regulatory subunit cyclin B1 (Fig. 1B). Several signaling pathways interact to ensure the maintenance of low MPF activity in the prophase I-arrested oocyte and then to activate MPF when it is time to reenter meiosis.

Maintaining a High cAMP Level in the GV Oocyte A high level of cyclic adenosine 30 ,50 -monophosphate (cAMP) in the oocyte is responsible for preventing CDK1 activation and for maintaining oocyte arrest at prophase I (Fig. 2A). The spontaneous meiotic maturation of mouse oocytes is prevented when they are cultured in the presence of the cAMP analog dibutyryl cAMP (dbcAMP) or cAMP phosphodiesterase (PDE) inhibitors such as isobutyl methyl xanthine (IBMX) and milrinone. PDE

inhibitors maintain high levels of oocyte cAMP by preventing its degradation. Accordingly, the oocytes from mutant mice lacking PDE3A are permanently arrested at the GV stage leading to female infertility [2]. Genetic studies in mouse support the hypothesis that cAMP synthesized by oocytes themselves is necessary to maintain their meiosis arrest at the GV stage. Mouse oocytes are equipped with all of the necessary molecular machinery for the synthesis of cAMP. Adenylyl cyclase (AC) catalyzes the formation of cAMP from ATP. Mouse oocytes that cannot express the AC3 isoform of AC can no longer be arrested at the GV stage in vivo [3]. Gs protein, which is stimulated by G protein-coupled receptor 3 (GPR3) present in the oocyte plasma membrane, has been shown to stimulate AC3. Therefore, blocking a subunit of Gs in follicle-enclosed mouse oocytes prevents cAMP synthesis and causes spontaneous meiosis resumption [4]. Similarly, GPR3-null mouse oocytes also precociously resume meiosis within the follicles in vivo [5]. Experimental evidence indicated that intrinsic cAMP production by oocyte is required but not sufficient to maintain the GV arrest. cAMP and cyclic guanosine

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FIG. 2 Mechanisms of meiotic prophase maintenance and gonadotropin-induced resumption of meiosis. (A) Prophase I-arrested oocytes in fully grown follicles require the synthesis and maintenance of high levels of cAMP, which is produced via the activation of the GPCR/Gs/AC pathway. A high cAMP level is maintained in the oocyte by preventing its degradation by inhibiting PDE3A. Mural granulosa cells (GCs) produce natriuretic peptide precursor type C (NPPC), which stimulates the generation of cyclic guanosine monophosphate (cGMP) by NPR2 (a guanylyl cyclase) present in cumulus GCs. cGMP enters the oocyte via Cx37 gap junctions and prevents PDE3A from hydrolyzing cAMP in the oocyte. cAMP activates protein kinase A (PKA) that in turn activates the Wee1B kinase and inhibits the CDC25B phosphatase leading to the inactivation of CDK1. Also shown in the figure is the constant degradation of cyclin B1 (cycB1) by the APCCDH1 that prevents MPF activation in prophase I-arrested oocytes. The APCCDH1 itself is inhibited by early mitotic inhibitor-1 (EMI1) in prophase I-arrested oocytes. Low MPF activity also favors the activation of protein phosphatase 1 (PP1) that constantly dephosphorylates meiotic proteins. (B) A preovulatory surge of LH causes closure of gap junctions throughout the follicle halting the supply of cGMP to the oocyte. This in turn increases cAMP hydrolysis by PDE3A. Low levels of cAMP and PKA can no longer activate WEE1B and inactivate CDC25B and CDK1 becomes dephosphorylated and catalytically active. Active CDK1-cycB1 complex phosphorylates and inactivates PP1 and this favors the maintenance of the phosphorylation status of other CDK1 substrates. Phosphorylation of lamin A/C causes the breakdown of the nuclear envelope. Active CDK1 also phosphorylates several other meiotic proteins that favor the occurrence of GVBD and meiotic progression. P-phosphorylation.

monophosphate (cGMP) signaling cooperate in the maintenance of the high level of cAMP that is essential for oocyte meiotic arrest. cGMP is produced by the surrounding follicular cells and it passes into the oocyte through gap junctions where it inhibits cAMP hydrolysis by PDE3A [6]. LH causes closure of gap junctions throughout the follicle halting the supply of cGMP to the oocyte. This in turn increases the cAMP-hydrolytic activity of PDE3A and the resumption of meiosis [6]. Thus, the control of cGMP synthesis by the granulosa

cells (GCs) might be essential for maintaining meiotic arrest in fully grown oocytes. Mural GCs of mouse follicles express a natriuretic peptide precursor type C (NPPC) and the cumulus GCs surrounding the oocyte express NPPC receptor 2 (NPR2), which is a guanylyl cyclase. NPPC peptide can inhibit the spontaneous GVBD in cumulus cell-enclosed oocytes but not in denuded mouse oocytes. In support of these in vitro findings, oocytes from Npr2 or Nppc-null mice also resume meiosis spontaneously in vivo [7]. These results show

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that cGMP signaling plays an important role in maintaining the high cAMP concentration in the oocyte during prophase I arrest (Fig. 2A).

Maintaining a Low MPF Activity by cAMP in the GV Oocyte The best known cellular function of cAMP is to activate protein kinase A (PKA) in response to the G-protein coupled receptor (GPCR)-mediated signal transduction. Microinjection of the catalytic subunit of PKA can maintain their prophase I arrest. Similarly, although mouse oocytes lacking PDE3A fail to undergo GVBD, inhibition of PKA signaling can induce GVBD in these oocytes. PKA, however, does not phosphorylate CDK1 directly but it regulates and balances the activities of the WEE1B kinase and CDC25 phosphatase in such a way that CDK1 remains inactive. WEElB kinase and CDC25 phosphatase are direct substrates of PKA in mouse oocytes. Following the preovulatory LH surge, or after release from the follicle, the reduction in cAMP ultimately leads to activation of CDK1 (Fig. 2). CDK1 is inactivated when it is phosphorylated on Thr14 and Tyr15 by the WEE1/MYT1 kinases (Fig. 1B). In mouse oocytes, WEE1B has been shown to be a key CDK1 inhibitory kinase that phosphorylates CDK1 at Tyr15 [8]. WEE1B mRNA is translated in GV-arrested oocytes and down regulation of its expression enhances early resumption of meiosis [8]. Inhibition of WEE1B also causes GVBD in PDE3A-null mouse oocytes that normally fail to undergo GVBD upon release from the follicle. This also indicates that Wee1B acts downstream of PKA in maintaining prophase I arrest [8]. PKA directly phosphorylates WEE1B at Ser15 and enhances its kinase activity. Reduction of the MYT1 kinase also enhances the early resumption of meiosis in mouse oocytes. When both WEE1B and MYT1 are down regulated, the effect on the resumption of meiosis is additive compared with the down regulation of WEE1B or MYT1 alone [8]. Three isoforms of the dual specificity phosphatase CDC25 (CDC25A, CDC25B, and CDC25C) function in the regulation of the mammalian cell cycle to reverse the inhibitory phosphorylation on CDK1 by the WEE1/ MYT1 kinases. In mouse oocytes, CDC25B is the isoform responsible for CDK1 activation and the resumption of meiosis [9]. During the meiotic arrest of mouse oocytes, PKA inactivates CDC25B by phosphorylating it on Ser321. When CDC25B is phosphorylated by PKA, it binds to 14-3-3 proteins and is transported to the cytoplasm away from the phosphorylated CDK1 in the nucleus [10].

Regulation of Cyclin B1 Levels In the GV stage-arrested mouse oocyte, an adequate amount of cyclin B1 is already present for resumption

of meiosis and no new protein synthesis is required. Oocyte maturation in cyclin B2-null mice remains unaffected and the mutant female mice remain fertile. Thus, under physiological conditions, cyclin B1 seems to be capable of compensating for the loss of cyclin B2 for MPF activation and oocyte meiotic maturation. Activation of CDK1 kinase by removing its inhibitory phosphorylation is sufficient to cause the resumption of meiosis. However, cyclin B1 must be constantly degraded by the anaphase promoting complex (APC) to maintain the GV stage arrest in the mouse oocytes [11]. If cyclin B1 is not continuously degraded during the GV stage arrest, its concentration increases leading to MPF activation and spontaneous resumption of meiosis. The APC is a multisubunit ubiquitin E3 ligase that tags its substrates by polyubiquitination, and these are then identified and degraded by the 26S proteasome. The two APC substrate adaptors, CDC20 and CDH1, are required for the substrate specificity and activation of the APC in oocyte meiosis. In mouse oocytes, CDH1 is required for the APCmediated cyclin B1 destruction to arrest oocytes at prophase I. When CDH1 function is lost, mouse oocytes can no longer maintain arrest at prophase I even if they are cultured in medium that normally maintains prophase I arrest [11]. In the GV stage mouse oocytes, the APCCDH1 activity itself is inhibited by early mitotic inhibitor-1 (EMI1). Reduction of EMI1 in the GV stage mouse oocytes by injection of morpholinos delays GVBD by preventing the accumulation of cyclin B1 whereas EMI1 overexpression leads to cyclin B1 accumulation and GVBD [12]. Thus, besides the inactivation of CDK1 by phosphorylation, destruction of cyclin B1 also keeps MPF inactive during the meiotic arrest in oocytes [13] (Fig. 2A).

Balancing Protein Phosphatases Protein phosphatases (PPs) that dephosphorylate the CDK1 substrates have been found to be important in regulating both mitosis and meiosis. Accumulating evidence shows that concomitant inhibition of CDK1-opposing PPs is required together with the activation of CDK1 kinase during oocyte meiotic maturation. The GV stage mouse oocytes contain both PP1 and PP2A, and PP1 is localized to the nucleus and PP2A is localized to the cytoplasm. CDK1 inactivates PP1 in somatic cells by phosphorylating it on Thr320. PP1 phosphorylation is also increased following GVBD in mouse oocytes, and this phosphorylation is sensitive to roscovitine. PP2A consists of a scaffold A subunit, a catalytic C subunit, and a regulatory B subunit, and is a known cell cycle regulator in both somatic cells and oocytes. Among the three PP2A subunits, PP2A-A protein levels were

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remarkably reduced during oocyte maturation and fertilization, whereas PP2A-B and -C subunits’ expression was stable. Increased PP2A activity inhibited oocyte GVBD by counteracting CDK1 activation [14]. In contrast, genetically deleting Ppp2r1a in oocytes, a gene encoding for the major isoform of the PP2A-A subunit, facilitated GVBD, which suggested that PP2A functioned as an inhibitor of meiotic resumption [15]. In addition, PP2A is required to prevent precocious segregation of sister chromatids during oocyte meiosis I [14]. During meiotic division I, sister chromatids are attached to one another by cohesin complexes when homologous chromosomes form tetrads. During the MI-to-AI transition, cohesins on chromosome arms are phosphorylated and then cleaved by separase, while centromeric cohesins are protected by a PP2A-shugoshin complex through dephosphorylation [16–18]. E3 ubiquitin ligase CRL4 controls oocyte meiotic maturation by proteasomal degradation of protein phosphatase 2A (PP2A) scaffold subunit, PP2A-A. CRL4 core components include cullin 4A or B as a scaffold, damaged DNA-binding protein 1 (DDB1) as a linker, and the ring finger protein ROC1/2 (Fig. 3). By selectively forming protein complexes with its more than 90 substrate adaptors known as DDB1-CUL4-associated factors (DCAFs), CRL4 regulates a wide range of cellular processes. Compelling evidence indicates that CRL4DCAF1 is a crucial regulator of meiotic maturation in fully grown oocytes: oocyte-specific deletion of DDB1 or DCAF1 results in delayed meiotic resumption and female infertility. Although these knockout mice could ovulate in response to exogenous gonadotropin when they were young, most of their ovulated oocytes had abnormal morphologies, as characterized by the presence of GVs and the absence of PB1s [14]. The PP2A-A subunit is accumulated in DDB1-

FIG. 3 Schematic showing CRL4DCAF1 function for regulating PP2AA degradation to facilitate oocyte meiotic progression. Green, gray, and red lines indicated activity changes of PP2A, CRL4, and MPF during oocyte meiotic maturation, respectively [14].

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or DCAF1-deleted oocytes, whereas Ppp2r1a knockout in DDB1-deleted oocytes rescued most of the meiosis defects. Biochemistry assays show that CRL4DCAF1 binds to PP2A-A and targets it for poly-ubiquitination and proteasome degradation. These results demonstrate that CRL4DCAF1 ubiquitin E3 ligase is essential for promoting meiotic cell cycle progression by targeting PP2A for degradation in mouse oocytes (Fig. 3).

Oocyte Meiotic Resumption Although fully grown oocytes are able to spontaneously resume meiosis in vitro, physiological meiotic maturation is triggered by LH in vivo. A surge of LH from the pituitary gland triggers ovulation, oocyte maturation, and luteinization for successful reproduction in mammals. This event, an essential prelude to fertilization, has long piqued curiosity and is a focus of research in reproduction. Mitogen-activated protein kinase (MAPK) is a family of serine/threonine protein kinases that are widely distributed in eukaryotic cells. The extracellular signalregulated kinase-1 and -2 (ERK1 and ERK2), two best studied members of the MAPK family, are activated by an LH surge in the GCs of preovulatory follicles. In mammals, ERK1/2 activation in granulosa and cumulus cells is necessary for gonadotropin-induced oocyte meiotic resumption, while ERK1/2 activities in oocyte itself are not required for its spontaneous meiotic resumption in vitro [19]. To examine how MAPK signaling pathways in the GCs mediate the LH actions and provoke oocyte maturation in vivo, Fan et al. developed a mouse model that specifically did not produce ERK1 or ERK2 in the GCs. The sexually mature mutant female mice did not ovulate and were completely infertile. The ovaries of these mice contained preovulatory follicles but not corpora lutea; accordingly, concentrations of serum estradiol in the mice were elevated, whereas progesterone concentrations remained low. The lack of response to LH was also observed in sexually immature mutant mice treated with exogenous hormones; their oocytes remained meiotically arrested, and the cumulus oophorus did not expand. Neither luteinization nor follicle rupture occurred in these animals. This study defines ERK1 and ERK2 as master regulators of fertility that mediate the effect of LH on all components of the ovulatory response: oocyte maturation, cumulus expansion, luteinization, and follicle rupture. This study, together with others, suggests that upon LH surge, ERK1 and ERK2 phosphorylate the gapjunction protein connexcin 43 in the granulosa and cumulus cells, leading to a diminution in the flow of molecules, such as cAMP and cGMP, both between these cells and

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FIG. 4 LH induces oocyte meiotic maturation through the extracellular signal-regulated kinases 1 and 2 (ERK1 and ERK2). ERK1 and ERK2, in response to follicular exposure to LH, phosphorylate the gap-junction proteins in the granulosa and cumulus cells, leading to a diminution in the flow of molecules, such as cyclic AMP (cAMP), both between these cells and between the cumulus cells and the oocyte. In the oocyte, cAMP represses the resumption of meiosis and thus maturation; a reduction of cAMP in the oocyte (brought about through diminished contact between the oocyte and cumulus cells) results in a resumption of meiosis and thus oocyte maturation. The expansion of cumulus cells around the oocyte also depends on ERK1 and ERK2. These proteins also mediate LH-induced rupture of the follicular wall. [20].

between the cumulus cells and the oocyte. A reduction of cGMP and cAMP in the oocyte (brought about through diminished contact between the oocyte and cumulus cells) results in PKA inactivation. As a result, CDC25B cannot be phosphorylated and inactivated by PKA and this active CDC25B can enhance CDK1 activity by removing the inhibitory phosphorylation caused by WEE1B/ MYT1 kinase (Fig. 2B). The activation of MPF leads to a resumption of meiosis and thus GVBD (Fig. 4).

Completion of the First Meiotic Division After meiotic resumption, MPF activity continues to rise during the first meiotic division, declines with first PB extrusion, and abruptly returns as the egg rearrests at MII (Fig. 3). The decrease in MPF activity as oocytes exit meiosis I is necessary to allow for segregation of homologous chromosomes, in an analogous manner to that in mitosis and again is driven by cyclin B1 degradation [21]. Degradation of securin, an inhibitor of separase activity (see below), is also required for separation of homologs, in that a nondegradable securin construct prevents first PB extrusion and homolog disjunction [21]. Therefore, both cyclin B1 and securin need to be efficiently degraded during first meiosis, and a failure to degrade either protein leads to an arrest at MI.

Meiosis I in mammals requires both the APC and the spindle assembly checkpoint (SAC) to correctly segregate homologs. The APC is a large multisubunit protein complex that earmarks cyclin B1, securin, and other substrates for proteolysis by polyubiquitination. The tagging of a substrate with ubiquitin causes its recognition by the 26S proteasome, which immediately proteolytically cleaves the protein. The APC shows little activity toward substrates without binding one of two protein cofactors CDC20 and CDH1. The APCCDC20 activity is often limited to the metaphase-anaphase transition: primarily because CDC20 needs CDK1 phosphorylation to associate with the APC, and CDK1 activity is normally confined to the period in M-phase leading to anaphase [13]. Cyclin B1 and securin are not the only APC substrates. In fact, the APCCDC20 and APCCDH1 target a large range of substrates for degradation during mitosis. These substrates contain discrete destruction motifs that target them for degradation. The most wellcharacterized motif is the D-box, which is recognized by both the APCCDC20 and APCCDH1, and the KENbox, recognized by the APCCDH1 only. The SAC prevents premature activation of the APC by recruiting components of the mitotic arrest deficient (MAD) and budding uninhibited by benzimidazole (BUB) families and MPS1 to kinetochores that are either

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unattached or lack tension [22,23]. By preventing premature APC activation, the SAC allows full congression and alignment of sister chromatids on the metaphase spindle before allowing anaphase. In this way, daughter cells receive one of each pair of homologous chromosomes (in meiosis I) or sister chromatids (in meiosis II and mitosis). Activation of the SAC normally occurs during the period of prometaphase I and that an inactive SAC leads to gross errors in segregation during meiosis I. The main SAC proteins include MAD1, MAD2, BubR1 (MAD3), BUB1, BUB3, and MPS1. MAD1 regulates the chromosome congression during oocyte maturation; MAD2 affects the chromosome segregation in meiosis I and meiosis II; BUB1 prevents chromosome segregation errors and aneuploidy in meiosis I; BUB3 is required for correct separation of both the homologous chromosomes and sister chromatids in meiosis I and meiosis II, respectively; BubR1 regulates the chromosome alignment and meiotic progression in oocyte maturation; and MPS1 is required for the meiotic progression and chromosome segregation in mouse oocytes [24].

Homologous Chromosome Separation Meiotic resumption is followed by chromosome alignment and spindle organization at prometaphase I (ProMI). During meiotic division I, sister chromatids are attached to one another by cohesin complexes when homologous chromosomes form tetrads. The cohesin complex in mammalian oocytes is consisted by the meiosis-specific subunits REC8, SA3, SMC1β, and SMC3 (Fig. 5A). During the MI-to-AI transition, accurate homologous chromosome segregation is achieved by cohesion removal from chromosome arms, after all chromosome pairs are aligned along the spindle equatorial plane. During the metaphase-anaphase transition, cohesin’s REC8 subunit is cleaved by separase, which is a cysteine protease distantly related to the caspases. Separase is kept inactive for most of the cell cycle by binding to an inhibitory chaperone called securin. Securin is only removed at the metaphase-anaphase transition by the APC-mediated proteolysis (Fig. 5B) [25]. In contract, the sister chromatid cohesins persist at centromeres and hold the sister chromatids together until anaphase occurs in meiosis II. The mechanisms underlying the protection of centromeric cohesion from separase cleavage when resolution of chiasmata occurs during meiosis I are emerging. REC8 subunit of cohesin on chromosome arms is phosphorylated and then cleaved by separase, while centromeric cohesins are protected by a PP2A-shugoshin complex through dephosphorylation [16–18]. The shugoshin 1 and shugoshin 2 proteins localize at the centromeres to protect REC8 from cleavage and prevent precocious sister chromatid separation in meiosis I [24]. The function of shugoshins in meiosis I is achieved

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by recruiting the PP2A to centromeres, and PP2A causes dephosphorylation of REC8 to counteract its cleavagedependent phosphorylation (Fig. 5B). DNA topoisomerase II (TOP2) is an enzyme that resolves DNA topological problems by introducing transient double-strand breaks (DSBs). TOP2 activity introduces a DSB, passes an unbroken strand through this transient break, and then reseals this break. During mitosis, DNA topoisomerase II (TOP2) is required for sister chromatid separation. When TOP2 activity is inhibited, a decatenation checkpoint is activated by entangled chromatin. In contrast, an effective decatenation checkpoint did not exist in fully grown oocytes, as oocytes underwent the G2/M transition and reinitiated meiosis even when TOP2 activity was inhibited [26]. However, oocytes treated with the TOP2 inhibitor ICRF-193 had severe defects in chromosome condensation and homologous chromosome separation [26]. Furthermore, condensed chromosomes failed to maintain their normal configurations in matured oocytes that were treated with ICRF-193. On the other hand, sister chromatid separation and subsequent chromosome decondensation during the exit from meiosis were not blocked by TOP2 inhibitors [26]. These results indicated that TOP2 had a specific, crucial function in meiosis I. Subsequently, oocytes extrude a first PB (PB1) and are arrested at metaphase II (MII) to await fertilization. Dysregulated meiotic progression results in aneuploidy and embryonic development abnormality, which contributes to early abortion and female infertility [27].

Interkinesis Meiosis is uniquely characterized by two rounds of chromosome segregation that follow DNA replication, allowing the production of progeny cells that are haploid. These two rounds of meiosis are not interrupted by interphase. Specifically, after PB1 formation DNA replication does not occur, the nuclear envelope does not form and chromosomes do not decondense. The period in between the two meiotic divisions is defined as interkinesis. During interkinesis, most cyclin B1 is degraded, MPF levels drop to low levels and in some instances, it is at the same level as in a GV stage oocyte [28], however, there is no chromatin decondensation at this time. It is thought that ERK1 and ERK2, whose activities are high in meiosis I oocytes, substitute for MPF activity during interkinesis. ERK1 and ERK2 recognize a similar peptide sequence to MPF and so may share substrates. The activity of ERK1 and ERK2 is elevated in association with meiosis reinitiation, and plays pivotal roles in regulating oocyte meiotic cell cycle progression (for reviews, see [29,30]). After GVBD, ERK1 and ERK2 are activated in maturing oocytes, and involved in the regulation of microtubule organization and meiotic spindle

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FIG. 5 Chromosome cohesion and separation during oocyte meiotic maturation. (A) The architecture of the cohesin complex in oocytes resembles a ring consisting of SMC1, SMC3β, SA3, and REC8. SMC1 and SMC3β form long, intramolecular coiled-coil stretches and dimerize. The ends of the V-shaped SMC1/SMC3β heterodimer are linked by REC8, which thereby closes the ring. Cohesin holds DNA strands together by entrapping them in the center of the ring. Cleavage of REC8 by separase leads to opening of the ring and dissociation of cohesin from chromosomes, thus allowing them to be pulled to opposite poles in anaphase. (B) Homologous chromosomes are held together by the cohesin complex from their generation during last replication before meiosis entry. During meiotic maturation, lagging chromosomes with unattached kinetochores block the initiation of anaphase by inhibiting APCCDC20. Once all chromosomes have bioriented, the APCCDC20 is activated and induces the degradation of securin, thus liberating separase from its inhibitor. Activated separase cleaves REC8, leading to cohesin’s dissociation from chromosomes, segregation of chromatids to opposite poles, and the initiation of anaphase. The remaining cohesins at centromeric regions are protected from separase cleavage by PP2A/ Sgo complex, and hold the sister chromatids together during anaphase I and MII.

assembly [31]. Most importantly, ERK1/2 activities are essential for maintenance of metaphase II (MII) arrest. The immediate regulator of MAPK is MEK, which in oocytes is activated by MOS, a distinct MEK kinase that is expressed exclusively in germ cells. In Xenopus and mouse oocytes, MOS is required for activation of MAPK and MPF at reinitiation of meiosis, for the suppression of DNA replication during interkinesis and for maintenance of the second meiotic arrest. When Mos, or Erk1 and Erk2 themselves, are knocked out, oocytes partially enter an interphase stage immediately following first PB extrusion: the chromosomes decondensed after PB1 emission [32]. In addition, regulation of MPF reactivation in interkinesis in Xenopus and mammalian oocytes is attributed to the intrinsic ERK1/2 activity as well. Prompt MPF reactivation secures the oocyte from entering interphase. Under conditions that prevented MPF reactivation, the

oocytes decondensed their chromosomes, formed a nucleus, and entered interphase [28].

Maintenance of MII Arrest Having extruded the first PB, the oocyte must re-establish a high MPF level and remain arrested at MII until fertilized. In addition to preventing chromosome decondensation during interkinesis, MAPK cascade is the master regulator that maintains MII arrest. Oocytes have been ascribed an activity, cytostatic factor (CSF) that confers an ability to arrest chromatids at MII, although its exact composition is still debated. MAPK cascade is responsible for CSF activity in both frog and mouse eggs. MOS injection into frog and mouse embryos induces a metaphase arrest and that removal of MOS from frog or mouse eggs prevents a MII arrest.

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FIG. 6 A summary of ERK1/2 functions in mouse oocytes and zygotes. In the GV oocytes, endogenous ERK1/2 is not required for meiotic resumption (GVBD) and spindle assembly at MI. However, Erk1/2oo / oocytes showed abnormalities at anaphase I (AI): homologous chromosome migrations to the spindle poles were asynchronous, followed by delayed PB1 emission. Although Erk1/2oo / oocytes are able to develop to MII, the spindle formation and chromosome alignment were abnormal. These oocytes failed to be arrested at MII. They released the second polar bodies (PB2s) shortly after ovulation, and were arrested at another metaphase called “MIII.” Furthermore, zygotes derived from ERK1/2-deleted oocytes showed defects in the male pronucleus formation, indicating that ERK1/2 is required for efficient oocyte-embryo transition [32].

Male PN:

Tubulin:

Female PN:

Chromosome:

Normal development:

GV

Maturation

GVBD

MI

Fertilization

AI

MII

Zygotic genome activation

Zygote

With ERK1/2 depletion:

Degeneration

Not affected Not affected Disorganized chromosome separation

Most knowledge about the in vivo function of ERK1/2 cascade in mammalian oocytes was obtained from analyzing the phenotypes of Mos knockout mice. Mos knockout female mice are less fertile and MII oocytes derived from them were parthenogenetically activated without fertilization [33,34]. Phenotypically, the first meiotic division of Mos/ oocytes frequently produces an abnormally large PB [35,36]. In these oocytes, the spindle shape is altered and the spindle fails to translocate to the cortex, leading to the establishment of an altered cleavage plane. Based on these observations, people concluded that ERK1/2 were not necessary for GVBD and PB1 emission but were required for normal spindle assembly. However, not all ERK1/2 functions in oocyte development can be revealed by analyzing MOS-deficient oocytes. Particularly, Mos mRNA is only accumulated in the fully grown GV oocytes, and translated into proteins after GVBD. Therefore, it remains unclear if ERK1/2 were activated by other upstream signals (such as oocyte membrane receptors) and involved in early events of oocyte development. Zhang et al. knocked out Erk1 and Erk2 in mouse oocytes as early as the primordial follicle stage using the well-characterized Gdf9-Cre mouse model, and for the first time directly investigated the in vivo function of ERK1/2 in mouse oocytes [32]. In this novel mouse model, they observed that ERK1/2 activities in oocyte are dispensable for primordial follicle maintenance, activation, and follicle growth. Different from the Mos null oocytes, the ERK1/2-deleted oocytes did not undergo a full parthenogenetic activation characterized for pronuclear formation. However, the ovulated ERK1/2-deleted oocytes have poorly assembled MII spindles, spontaneously released PB2s, and were arrested at another metaphase called metaphase III (MIII) (Fig. 6). In addition,

2-cell

Emit PB2; Enter MIII

Male PN formation defects

ERK1/2-deletion prevented male pronuclear formation after fertilization, and caused female infertility. These results indicate that ERK1/2 activities are required for not only MII-arrest maintenance, but also efficient pronuclear formation in mouse oocytes (Fig. 6). The ultimate control point in the maintenance of metaphase arrest is in the prevention of cyclin B1 and securin degradation. Since both these proteins are degraded at anaphase-onset by polyubiquitination through CDC20-bound APC (APCCDC20) and the 26S proteasome there are at least three points of possible CSF-induced metaphase arrest. The first (A) is the most upstream control point and would control the level of cyclin B1 and securin synthesis. Therefore, MII arrest is maintained by increased synthesis of cyclin B1/securin. The second control point is at the level of the APC, either directly by negative regulation of the APC or indirectly by affecting the ability of CDC20 to switch on the APC. This is by far the most preferred mechanism. The third control point (C) is at the level of the 26S proteasome and here it is reduced proteasome activity that prevents degradation of polyubiquitinated cyclin B1/securin. So far, the potential regulation of the 26S proteasome in oocyte meiosis is pretty much a hypothesis awaiting experimental evidence [13].

OOCYTE CYTOPLASMIC MOLECULAR MATURATION Translational Activation of Dormant mRNAs In vertebrates, the GV stage-arrested fully grown oocytes contain a large amount of maternal mRNAs that are translationally dormant [37]. Upon meiotic

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maturation, many maternal mRNAs are translationally activated [38,39]. These temporally translated proteins play key roles in meiotic spindle assembly, MII arrest maintenance, and mRNA clearance during maternalzygotic transition (MZT). The mechanisms of temporal and selective activation of maternal mRNAs are complicated and diverse among species [40]. Cytoplasmic polyadenylation of the 30 untranslated region (30 -UTR) is largely correlated with mRNA stability and translational activation of the mRNA, and plays an essential role in oocyte maturation. Studies of mouse and Xenopus oocytes revealed that mRNA translational repression is partially accomplished by deadenylation and loop formation between mRNA termini preventing interaction of translation initiation factors and transcript degradation, allowing for longterm storage of mRNA. The stored transcripts are then recruited for translation in a spatiotemporal manner to drive essential processes during oocyte maturation when the oocytes are transcriptionally quiescent. Cytoplasmic polyadenylation of maternal mRNAs requires a cytoplasmic polyadenylation element (CPE) that binds specific trans-acting proteins [41]. Studies in Xenopus and mouse oocytes found that the CPE-binding protein-1 (CPEB1) mediates cytoplasmic polyadenylation of many CPE-containing mRNAs [42]. During Xenopus and mouse oocyte maturation, ERK1/2-triggered phosphorylation of CPEB on several serine/threonine residues is required for early activation of a class of maternal mRNAs, while a large fraction (70%–90%) of CPEB1 proteins undergo a polyubiquitination-dependent degradation in meiosis I [43,44]. This degradation causes a change in the CPEB/CPE ratio and results in activation of another class of mRNAs. By activating CPEB1, ERK1/2 couples the translation of a series of maternal mRNAs, including Dazl, Tpx2, and Btg4, to meiotic maturation. The translation products of these maternal mRNAs are required for meiotic divisions and MZT (Fig. 7). Nucleoside analogs, 30 -deoxyadenosine (30 dA) function as inhibitors of RNA elongation once incorporated by terminating extension. in vitro meiotic maturation of murine, bovine, and porcine oocytes was blocked by 30 dA treatment. In addition, female knockout and conditional siRNA knockdown of Cpeb1 resulted in lack of fertile gametes and severe ovarian abnormalities in mouse, respectively [37,45]. Furthermore, knockout of another cytoplasmic polyadenylation-associated gene in female mice, poly(A) binding protein cytoplasmic 1-like (Pabpc1l, also known as embryonic poly(A) binding protein, ePAB), prevented oocyte maturation in vitro and in vivo precluding formation of embryos whereas male knockouts were fertile [46]. These findings suggest that cytoplasmic polyadenylation is necessary for completion of both components of oocyte maturation, as successful nuclear maturation is not an indicator of the molecular

maturation status of the oocyte, which may affect downstream developmental stages. Although the elongation of poly(A) tails in cytoplasm is essential for oogenesis, the poly(A) polymerases responsible for the cytoplasmic polyadenylation in mammalian oocytes have not been identified. Several poly(A) polymerases responsible for the cytoplasmic polyadenylation are identified in yeast (Cid1), Caenorhabditis elegans (Gld-2), and Xenopus (xGld-2). Poly(A) polymerase D4 (PAPD4) is a mouse homolog of Xenopus xGLD-2. PAPD4 associates with cytoplasmic polyadenylation components, CPEB and cleavage and polyadenylation-specific factor (CPSF) described in Xenopus oocytes. In spite of the ubiquitous expression of PAPD4 in mouse tissues, the Papd4 knockout mice were normal and healthy. Moreover, Papd4 disruption did not affect the poly(A) tail elongation in oocytes using reporter RNAs [47]. Thus, the other PAPD family members, such as PAPD5 and PAPD7, may play a redundant role with PAPD4 [48].

Maternal mRNA Decay This transition from a maternal to a zygotic mode of development is called the MZT. Maternal mRNAs, which are synthesized and stored during oocyte growth, serve as the maternal contribution that supports early embryo development but undergo general decay after meiotic resumption. In mammals, oocyte meiotic maturation is a prolog to MZT, which triggers a transition from mRNA stability to instability, suggesting the existence of an active mechanism that triggers mRNA decay in oocytes [49]. About 90% of maternal mRNAs are degraded by the two-cell stage in mouse embryos [50]. The destruction of transcripts during the GV to MII transition is selective rather than promiscuous in mouse oocytes [51]. Particularly, transcripts associated with meiotic arrest and the progression of oocyte maturation, such as oxidative phosphorylation, energy production, and protein synthesis, were dramatically degraded. Selective degradation of these maternal transcripts is a prerequisite for transition from oocyte meiosis to blastomere mitosis, as well as transcriptional activation of the zygotic genome [45,52]. Aberrant degradation or maintenance of certain classes of transcripts during oocyte maturation could be deleterious to oocyte quality and affect developmental competence, as demonstrated in Btg4 knockout oocytes (see below).

BTG4 and CCR4-NOT RNA Deadenylase The mammalian B-cell translocation gene (BTG) family comprises six proteins (BTG1, BTG2, BTG3, BTG4, TOB1, and TOB2), which regulate cell cycle progression in a variety of cell types. They are characterized by the

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FIG. 7 A diagram showing role of ERK1/2 in triggering maternal mRNA translation and degradation during mouse oocyte maturation and maternal-zygotic transition (MZT). Upon oocyte meiotic resumption, ERK1/2 is activated by upstream kinases and triggers CPRB1 phosphorylation and CRL1βTrCP-dependent degradation. The phosphorylation and partial degradation of CPEB1 stimulate polyadenylation and translational activation of maternal mRNAs including Dazl and Btg4. The accumulated DAZL proteins lead to further translational activation of maternal mRNAs such as Tpx2 and Btg4, whereas BTG4 and CNOT7 target polyadenylated maternal mRNAs to degradation. Btg4 mRNAs are stored in fully grown oocytes. During meiotic resumption, the ERK1/2 cascade and CPEB1 are sequentially activated. CPEB1 stimulated polyadenylation and translation of Btg4 mRNA by interacting with three cytoplasmic polyadenylation elements (CPEs) in its 30 -UTR. In maturing oocytes, BTG4 proteins mediate the interaction between CNOT7 or 8 and eIF4E, and recruit the CCR4-NOT deadenylases to the translating maternal transcripts. As a result, maternal transcripts are degraded during oocyte maturation and fertilization, which is a prerequisite for zygotic genome activation. By initiating these hierarchical maternal mRNA regulation processes, ERK1/2 functions as a meiotic cell cycle-coupled licensing factor of maternal mRNA translation. Red and green curves represent maternal and zygotic transcripts, respectively. The gray shadow represents the time frame of BTG4 protein expression [31,45]

conserved N-terminal BTG domain that spans 104–106 amino acids. The conserved BTG domain is a proteinprotein interaction module, which is capable of binding to CNOT7 and CNOT8, two deadenylase subunits of the CCR4-NOT deadenylase complex. Among the BTG family proteins, BTG4 is an MZT licensing factor in mouse. BTG4 bridges CCR4-NOT deadenylase to eIF4E, a key translation initiation factor, and plays a permissive role in maternal mRNA decay. While the knockout of genes encoding other BTG family proteins only causes minor developmental defects in moues, Btg4 null females produce morphologically normal oocytes but are infertile due to early developmental

arrest [45]. A RNA sequencing study indicated that BTG4 is crucial for maternal mRNA clearance. A large proportion of maternal transcripts were degraded during oocyte maturation and after fertilization, but they were not degraded in oocytes and zygotes of Btg4 null females. Mechanistically, BTG4 triggered decay of maternal mRNAs by mediating the CCR4-NOT-dependent deadenylation of their poly(A) tails (Fig. 7). Importantly, translation of maternal Btg4 and Cnot7 mRNAs is coupled with meiotic maturation. Although Btg4 and Cnot7 mRNA levels were high in the GV oocytes, expression of their proteins was only detected after GVBD, reached the maximal level at the MII stage,

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and quickly decreased after the two-cell stage. The 30 -UTR of mammalian Btg4 mRNAs contains three putative CPEs which are crucial for translation. During meiotic maturation, oocyte intrinsic MAPK cascade and CPEB1 trigger polyadenylation and translation of Btg4 mRNA stored in fully grown oocytes by targeting these CPEs. The accumulated BTG4 and CNOT7 proteins in turn, mediate maternal mRNA deadenylation and degradation [45,53]. U0126, the inhibitor of ERK1/2 activation, blocked BTG4 accumulation during oocyte meiotic maturation. In oocyte-specific Erk1/2 knockout mice, BTG4 protein remained undetectable in oocytes at the MII stage, and maternal mRNA decay was blocked. Therefore, a negative feedback regulation mechanism is formed to ensure transient, but not prolonged translation of proteins crucial for oocytes and zygotes. This is a key step in oocyte cytoplasmic maturation that determines the developmental potential of mammalian embryos. The function of ERK1/2 in regulating maternal mRNA translation and degradation during oocyte maturation and MZT is summarized in Fig. 7.

Zinc Finger Protein 36 Like 2 In Xenopus oocytes, a significant fraction of transcripts that are cytoplasmically polyadenylated at meiotic-phase transitions contain, in addition to CPEs, (A + U)-rich element (ARE) sequences (characteristic of mRNAs regulated by deadenylation). Among these is the mRNA encoding zinc finger protein 36 like 2 (ZFP36L2, also known as C3H-4 in Xenopus), a ARE-binding protein that accumulates after meiotic resumption [54]. ZFP36L2 belongs to a family of zinc finger proteins containing tandem zinc-binding motifs characterized by three cysteines followed by one histidine (CCCH). Through their zinc fingers, these proteins can bind to mRNAs containing ARE in their 30 -UTR. Meanwhile, ZFP36L2 recruits the CCR4-NOT deadenylase complex to ARE-containing mRNAs by direct binding with CNOT6, a catalytic subunit of CCR4-CNOT complex. In this way, ZFP36L2 triggers poly(A) tail shortening and decay of AREcontaining mRNAs. Depletion of ZFP36L2 in Xenopus oocytes prevents MI-MII transition. The role of ZFP36L2 in oocyte maturation is conserved in mammals. Zfp36l2 knockout female mice are infertile. The number of oocytes being ovulated by these mice is reduced, and embryos derived from these oocytes showed developmental arrest at two-cell stage [55]. These results indicated that ZFP36L2 is essential for oocyte cytoplasmic maturation in vivo. The cooperative activities of the CPEs and the AREs define the precise activation times of the mRNAs. An “early” wave of cytoplasmic polyadenylation activates a negative feedback loop by activating the synthesis of

ZFP36L2, which in turn would recruit the deadenylase complex to mRNAs containing both CPEs and AREs. This negative feedback loop is required to exit from metaphase into interkinesis and for meiotic progression. Sequential waves of polyadenylation and deadenylation ensure irreversible, self-sustained, meiotic phase transitions by controlling discrete states of MPF, APC, and CCR4-NOT activities. The multiple translationdependent positive and negative feedback loops help to keep the oocyte from slipping rapidly back and forth between cell cycle phases.

Zygotic Arrest-1 (ZAR1) and -2 (ZAR2) Zar1 is the first maternal effect gene identified in mouse. Zar1 null female mice generate fully grown oocytes and the eggs can be fertilized, but the resulting embryos fail to develop beyond two-cell stage. On the other hand, recent results in zebra fish indicated that Zar1 is also critically required for early oogenesis, by repressing excessive mRNA translation thereby preventing overload of maternal proteins in early oocytes [56]. Loss of Zar1 causes early oogenesis arrest and female-to-male sex reversal in this species. In addition, a Zar1-like gene that is predominantly expressed in oocytes and zygotes were identified in Xenopus and mouse [57,58]. This Zar1 homolog was designated as Zar2 or Zar1l, but the in vivo function of this oocyteenriched gene has not been investigated. The molecular regulatory mechanism of ZAR1 in oocytes is largely unknown. Both Zar1 and Zar2 are conserved in vertebrates and contain an atypical plant homeodomain (PHD) zinc finger domain in C-terminus [59]. in vitro results suggest that its Xenopus homolog may function as a RNA-binding protein to regulate RNA translation [60]. Both ZAR1 and ZAR2 bind to translational control sequence (TCS) of Wee1 and Mos mRNAs and repress their translation in immature oocytes. In addition, ZAR1 was reported to be associated with known translation factors, such as CPEB and ePAB. Collectively, ZAR1 and ZAR2 may function as components of a maternal translational complex to recruit other translational regulators and repress mRNA translation in early oocytes. Future study will examine how the translational complex is regulated by ZAR1 and ZAR2 in mammalian oocytes.

OOCYTE CYTOPLASMIC ORGANELLE MATURATION Mitochondrial Number and Distribution Mitochondria contain one or more copies of their own genome (mtDNA), which encodes a total of 13 genes with

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critical functions in cellular metabolism. They are the most important cellular organelle in cytoplasm, and their variation in distribution, their capacity of generating ATP as well as their DNA copy number play an important role in oocyte maturation and early embryo development via their role in cell survival and apoptosis. It is accepted that, apart from a very short period immediately after fertilization, mtDNA does not replicate until blastocyst implantation, and the development of fertilized eggs is dependent on the existing pool of mitochondria. During follicular growth, the number of oocyte mitochondria rises from  10,000 to  200,000 [61]. In the mouse, oocytes with as few as 4000 copies of mtDNA can be fertilized and develop to blastocyst stage, while the mature oocyte has a critical postimplantation developmental threshold of 40,000–50,000 copies of mtDNA [62]. An average of 256,000  213,000 mitochondrial genomes is found to exist in each human oocyte. The developmental potential of the embryo and the outcome of in vitro fertilization (IVF) have been also shown to be related to the ATP content of human oocytes. In unfertilized oocytes and those arrested at the pronuclear stage after fertilization, the mtDNA numbers were lower than in fertilized eggs and cleaved eggs, respectively. Not only the number of mitochondria, but also their distribution and activity affect oocyte maturation events and embryo development. Mitochondria are frequently organized in dynamic interconnected networks, and they move to areas of high-energy consumption [63]. When the oocytes were treated with the mitochondria-targeted compound carbonylcyanide-m-chlorophenylhydrazone (CCCP), which is a proton gradient uncoupler, their meiotic maturation was arrested [64]. In oocytes collected from small follicles, mitochondria were observed mainly in the cortex, while accumulation of mitochondria in the peripheral cytoplasm and around the GVs was characteristic of fully grown GV oocytes collected from large follicles. During oocyte maturation, numerous mitochondria distributed around the nuclear area, which provides energy for the GVBD, spindle organization, and chromosome segregation (Fig. 8A). Larger mitochondrial foci were also found to move to the inner cytoplasm in mature porcine oocytes [65]. Compromised oocyte quality is associated with aberrant mitochondrial rearrangement and low ATP levels [66]. Distribution of mitochondria in oocytes matured in vitro is slightly different from that of oocytes matured in vivo. Compared with the oocytes matured in vivo, in which large mitochondrial foci were distributed throughout the cytoplasm (Fig. 8B), mitochondria were not observed in the central cytoplasm in most pig oocytes matured in vitro (Fig. 8C) [65]. In human GV oocytes, most mitochondria localize predominantly in the noncortical region of the cytoplasm, while they occupy the entire area of the cytoplasm at GVBD [67]. The

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mitochondria were more abundant in the inner cytoplasm than in the peripheral region in most of the human oocytes matured in vivo [68]. These results may partially explain the reduced developmental potential of oocytes matured in vitro compared with those matured in vivo. Mitochondrial dynamics change mitochondrial morphological features and numbers as a part of adaptive cellular metabolism. Mitoguardin-1 and -2 (Miga1/2) encode to closely related mitochondrial outer-membrane proteins which promote mitochondrial fusion. Knockout mouse strains were generated for the mammalian Miga1 and Miga2. Miga1/2/ females show greatly reduced quality of oocytes and early embryos and are subfertile. Mitochondria became clustered in the cytoplasm of oocytes from the germinal-vesicle stage to meiosis II; production of reactive oxygen species (ROS) increased in mitochondria and caused damage to mitochondrial ultrastructures. In addition, reduced ATP production, a decreased mitochondrial-DNA copy number, and lower mitochondrial membrane potential were detected in Miga1/2/ oocytes during meiotic maturation. These changes resulted in low rates of polar-body extrusion during oocyte maturation and reduced developmental potential of the resulting early embryos. Vitamin C treatment remarkably reduced ROS levels and partially rescued the normal mitochondrial distribution in Miga1/2/ oocytes. Meanwhile, ATP supplementation also partially reversed the polar body emission (PBE) defects and reduced ROS levels in Miga1/2/ oocytes. Therefore, MIGA1/2-regulated mitochondrial dynamics are crucial for mitochondrial functions, ensure oocyte maturation, and maintain the developmental potential (Fig. 8A).

Cortical Granule Migration Cortical granules are membrane-bound organelles located in the cortex of unfertilized oocytes. They range in size from 300 to 400 nm in diameter in the human. Cortical granules undergo a substantial distribution change in oocyte cortex during meiotic maturation. In immature mouse GV oocytes, cortical granules exist as a continuous cortical layer with some interior granules. Mature oocytes have an asymmetric cortical distribution with a cortical granule-free domain (CGFD), overlying the metaphase II spindle where there is no microvilli distribution and where sperm does not penetrate during fertilization, occupying 40% of the cortex (Fig. 8D and E). The mean cortical granule densities of the entire cortex of mouse GV oocytes and the granule-occupied cortex of mature oocytes are 34 and 43 cortical granules/100 μm2, respectively. The mean total numbers of cortical granules/ oocyte are 4127 (mature) and 7440 (GV), respectively [69]. In pig oocytes, cortical granules distribute in the

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FIG. 8 Distribution of oocyte cytoplasmic organelles during oocyte maturation. (A) Mitochondrial aggregate around the nuclear area at the time of GVBD in oocytes. Deletion of Miga1/2 or CCCP treatment results in mitochondrial clustering, reduces ATP levels and mitochondrial DNA (mtDNA) copy numbers, and increases reactive oxygen species (ROS) levels in oocytes; these changes cause meiotic and developmental defects. Addition of vitamin C or ATP to the oocyte culture medium partially reverses these problems [64]. (B) and (C) Mitochondria distribution in pig oocytes matured in vivo (B) and in vitro (C). Arrow indicates the first polar body. Mitochondrial clusters fail to migrate to the central ooplasm in in vitro matured oocytes. (D) and (E) Distribution of cortical granules (CGs) in mature mouse oocytes. The CGs distribute beneath the oolemma, with a cortical granule-free domain (CGFD) overlying the meiosis II metaphase spindle apparatus.

cortex cytoplasm of oocytes at the GV stage with a mean number of 33.8  7.3 cortical granules/100 μm2 of cortex. The migration of cortical granules to the cortex continues during maturation, and cortical granules reach the cortex and form a continuous monolayer under the oolemma when maturation proceeds to metaphase I and metaphase II [70]. In the human, cortical granules synthesis is still observed at the GV stage, particularly at the onset of resumption of meiosis, while the other report indicates that there is no cortical granules synthesis at this stage, and the mean numbers of cortical granules per 10 μm of the oocyte linear surface are 8.0  2.22 and 8.1  1.38 in the GV oocytes and in vivo matured MII oocytes, respectively [71]. A well-defined F-actin band exists in the GV oocytes from small antral follicle, preventing the cortical granules from migrating to the periphery. This band gradually became disorganized at the GV stage as oocytes resumed meiosis, when cortical granules are apparently migrating to the surface. It is reported that cortical granules behave differently in mouse oocytes matured under different conditions, and cytoplasmic maturity is not fully achieved in the in vitro matured oocytes [72], which may hamper cortical granule exocytosis and developmental potential after fertilization.

Redistribution of ER and Golgi Complex The migration of the ER to the cortex during oocyte maturation is thought to play an important role in rendering the ER competent to generate the calcium transients after fertilization. Endoplasmic reticulum Ca2+ ([Ca2 + ]ER) increases, and alteration of intracellular Ca2+ ([Ca2 + ]i) homeostasis undermines maturation in mouse oocytes. The ability to mount the full complement of oscillations is only achieved at the end of oocyte maturation. Indeed, in vitro fertilized GV oocytes show fewer [Ca2+]i oscillations and each [Ca2+]i rise exhibits lesser duration and amplitude than those observed in fertilized MII eggs, and several parameters such as ER reorganization and increase in [Ca2+]ER) are thought to involve in this process. The sensitivity of Ca2+-releasing mechanism is also increased by increased reactivity of IP3R molecules, through which the ER responds to the generation of IP3 after fertilization. In human GV stage oocytes, ER is organized in a fine network extending throughout the cortex and the cell interior, while MII oocytes have large distinct ER clusters throughout the cortex and the cell interior [73]. During GV-MII transition, IP3R expression increases by 50%, while the ability to release Ca2+ in response to IP3 almost doubles [73]. Therefore, ER undergoes both distribution and constitution changes

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during oocyte maturation, which accounts for an increased ability to release Ca2+. Human MII oocytes matured in vitro fail to support such increases in IP3R abundance and Ca2+-release ability despite an apparently normal ER distribution pattern, suggesting that a reduced response to IP3 may explain the reportedly lower developmental competence of in vitro matured oocytes [73]. The GV oocytes have Golgi complexes distributed throughout the cytoplasm. Golgi complexes decrease at the edge of the oocyte and become fragmented with the progression of oocyte maturation [74]. Dispersion of mini-Golgis occurs in oocyte of various species. In mature oocyte, Golgi complex, lysosomes, and ribosomes are very rare.

chromosome movement, homologous chromosome separation, spindle anchorage, and vesicle organelle transport may require interactions between microfilaments and microtubules. For example, during oocyte maturation, the distribution of the ER undergoes major modifications guided by microtubules and microfilaments to make the oocyte more competent in the generation of intracellular Ca2+ oscillations that are pivotal for triggering egg activation [71]. MATER, a cortical protein complex which has close relationship with actin functions, is required for ER distribution and Ca2+ homeostasis in oocytes, likely due to its effect on lattice-mediated ER positioning and/or redistribution [78]. How the two systems of the cytoskeleton precisely interact to perform their functions remain little known.

Cytoskeleton Dynamics

Evaluation of Cytoplasmic Quality and Improvement of Cytoplasmic Maturation

Oocyte undergoes asymmetric meiotic division, and both chromosomes and organelles redistribute during oocyte maturation. The cytoskeletal microfilaments and microtubules present in the cytoplasm promote organelle movements and act on chromosome movement and segregation. Disruption of either microtubules or microfilament causes failure of chromosome movement and separation, and oocyte is arrested at metaphase stage. The cortical polarity establishment depends on F-actin which forms a dense layer beneath the oolemma, with an actin-cap formed above the meiotic apparatus, which is also required for polarity maintenance. In human, highresolution confocal microscopy demonstrates that in MII, but not GV stage, oocytes cortical actin is more abundant in the vicinity of the spindle. A cloud of dynamic actin filaments trail behind the chromosomes, generating actin flows and finally cytoplasmic streaming, to drive their peripheral movement [75]. At telophase, an F-actin ring is formed at the midbody which is required for final chromosome segregation. Microfilaments also control cortical granule redistribution, since its disruption influenced cortical granule migration. The actin nucleation factor actin-related protein 2/3 complex and its nucleationpromoting factors, formins and Spire, and regulators such as small GTPases, partitioning-defective/protein kinase C, Fyn, etc. are all involved in microfilament functions [76]. Microtubules control mitochondria redistribution. Disassembly of microtubules with nocodazole inhibits both mitochondrial aggregations to the GV area and their inward movement to the inner cytoplasm during oocyte maturation, as well as the translocation of mitochondria to the peri-pronuclear region during fertilization, whereas disruption of microfilaments by cytochalasin B has no effect in both human and pig oocytes [65,77]. The completion of several dynamic events, including

Traditional methods for the evaluation of oocyte quality are based on morphological characters of the follicle, cumulus-oocyte complex, PB, and/or meiotic spindle. Although the use of these morphological predictors to evaluate oocyte quality is controversial, such a grading system can provide valuable information for oocyte developmental competence. To judge oocyte cytoplamic maturation or quality, several markers (such as mitochondrial status, cortical granule distribution, GSH level, etc.) may be used as indicators of oocyte cytoplasmic maturation. However, these cellular predictors are invasive. Competent immature oocytes have finish their growth phase and show decreased glucose-6-phosphate dehydrogenase (G6PDH) activity. Brilliant cresol blue (BCB) is a dye that is degraded by G6PDH and its staining can be used to distinguish oocytes that have finished their growth phase (BCB +) from those that are still growing and are less competent (BCB). BCB staining of the GV oocytes has been used to evaluate oopalsmic quality in various species [79]. Clinically, the assessment of human oocyte cytoplasmic maturation has been hampered by the lack of reliable and nondestructive predictors of viability. During in vitro oocyte maturation, the spontaneous meiosis resumption may start at various times causing heterogeneity in the nuclear stage and also in cytoplasmic maturation or non-synchronization of the two events. The synchronization of nuclear progression in a population of oocytes or synchronization between nuclear and cytoplasmic maturation in an individual oocytes is important for improving oocyte developmental competence. This can be achieved by a short-term inhibition of meiotic resumption during the early phase of in vitro culture by increasing cAMP level and inhibiting MPF activity. Such pharmacological compounds can allow a prolonged oocyte maturation period is to promote a longer interaction between the

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immature oocyte with cumulus cells, and thus synchronization of nuclear and cytoplasmic maturation processes. The presence of PDE3-I, Org9935, or cilostamide has a beneficial effect on normal spindle and chromosome configurations, synchronization of nuclear and ooplasmic maturation of human oocytes, and thus subsequent embryonic development [80]. It is now well known that abnormal mitochondrial distribution, decreased number and mtDNA mutations/ deletions are related to increased abnormalities of the nuclear maturation (GVBD, spindle formation, chromosome segregation) and decreased embryo development potential of oocytes. Age-associated decline in female fertility is largely due to ooplasmic mitochondrial dysfunctions. Although oocyte has its surveillance mechanisms for eliminating mutated mtDNA to some extent, and thus preserving mitochondrial functions over the generations, aging-related mitochondrial mtDNA mutations and/or deletions do accumulate in the aged oocyte, which attributes to the deterioration of oocyte cytoplasm quality. Various reproductive technologies including transfers of heterologous ooplasm, GV, spindle, PB, and pronuclei have recently been tested in animals and human to improve developmental potential of oocytes or rescue mtDNA defects by partially or fully replacing mtDNA and thus reduce risks of motherto-child mtDNA disease transmission. Such manipulations can not only provide more and/or healthy mitochondria, but also supply other cytoplasmic components including other organelles, mRNAs and proteins, but mitochondrial heteroplasmy and tri-parental ethnic issue still cause concerns. Autologous cumulus cell or oogonial precursor cell mitochondria transfer is also reported, but the efficacy needs further validation.

OOCYTE EPIGENETIC MATURATION Epigenetics, the cell-type-specific and inheritable interpretation of genetic material, relies mainly on DNA and histone modifications to regulate gene transcription. Mammalian oocytes undergo more drastic epigenetic changes than do somatic cells during development. This coincides with their nature in preserving genetic and epigenetic information over an extended time span, and transferring them to the next generation [81–83]. Histone modifications and histone exchanges ensure the appropriate expression of genes involved in oogenesis in the absence of DNA replication, and poise the zygotic genome for transcriptional activation after fertilization.

Epigenetic Modifications in the GV Oocyte In the mouse ovary, the first wave of oocyte growth and differentiation is synchronous and is also the time

at which maternal-specific genomic imprints are established. This process of epigenetic modifications or “epigenetic maturation” is capable of affecting gene expression without a change in DNA sequence and ultimately confers the mammalian genome with a sexspecific mark or genomic imprint essential for embryonic development. Importantly, the oocyte genome is also subject to additional levels of regulation, and functional differentiation of large-scale chromatin structure provides an important epigenetic mechanism for the developmental control of global gene expression. For example, coincident with follicular activation, an oocyte-specific linker histone (H1Foo) is loaded into the mouse oocyte nucleus) consistent with a possible role for multiple subtypes of linker histone H1 during oogenesis. Moreover, dynamic changes in chromatin structure and function occur during oocyte growth. Morphological transitions in the GV were originally recognized in many mammalian species including human. Chromatin in growing mouse oocytes (Fig. 9A) is initially found decondensed in a configuration termed nonsurrounded nucleolus (NSN). With subsequent growth and differentiation, oocytes undergo a dramatic change in nuclear organization in which chromatin becomes progressively condensed (Fig. 9B), forming a heterochromatin rim in close apposition with the nucleolus, thus acquiring a configuration termed surrounded nucleolus (SN). Histone modifications, such as acetylation, methylation, and phosphorylation, play important roles in the regulation of chromatin structure and gene expression. In general, histone acetylation leads to the relaxation of chromatin structure and thus correlates with gene activation. In contrast, histone deacetylation leads to condensation of chromatin structure and thus correlates with gene repression. Therefore, the core histone tails of GV chromatin should become less acetylated as oocytes grow, because chromatin condenses and gene expression is silenced during oocyte growth. However, it was shown that fully grown mouse GV oocytes were fully acetylated at all the lysine residues on H3 and H4, but underwent deacetylation after GVBD and became acetylated again in one-cell embryos [84]. In fact, analysis revealed that the acetylation of H3K9, H3K18, H4K5, and H4K12 increased during mouse oocyte growth and that fully grown GV oocytes showed the most modifications [85]. Furthermore, the level of histone acetylation in mouse SN oocytes was shown either similar to or even higher than that in the NSN oocytes [85]. Similarly, the histone H3 acetylation status was maintained, while the chromatin configuration changed from decondensed to a perinucleolar heterochromatin sheath during the growth of the pig oocytes [86]. The inhibition of histone deacetylase (HDACs) with trichostatin A (TSA) inhibited the deacetylation of histone H3 and post-GVBD chromosome

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FIG. 9 Conformation and histone H3 methylation changes in the GV oocytes (A–B) DNA staining showing nonsurrounded nucleolus (NSN) and surrounded nucleolus (SN) chromatin configurations in GV of mouse oocytes. (C–D) Immunofluorescent staining of histone H3 lysine-4 trimethylation (H3K4me3) in mouse oocytes with NSN and SN-type nucleus. (E–F) Merged GV images of DNA and H3K4me3 staining.

condensation in pig oocytes [86]. Furthermore, although inhibition of HDACs with TSA-induced chromatin decondensation, it did not restore transcriptional activity in mouse SN oocytes [84]. Therefore, histone acetylation may not be associated with transcriptional activation while deacetylation may not be associated with chromatin condensation during oocyte growth.

Exchanges of Histone Variants During Oocyte Maturation Histones are the main structural proteins that package eukaryotic DNA into chromatin. Changes in nucleosome composition that occur as a result of the replacement of canonical histones with their variants also play important roles in chromatin remodeling. A variety of variants for histones H2A, H2B, and H3 have been identified in eukaryotes. Among these, histone H2A has the largest number of variants, including H2A.Z, H2A.X, macroH2A, and H2A.Bbd, which are distinguished from the canonical H2A by variations in the length and sequence of their C-terminal tails. Turnover and exchange of histone variants actively occurs during mouse oogenesis, even in the absence of transcription. Postnatal mammalian oocytes undergo several developmental transitions in the absence of DNA replication,

thus making oogenesis an ideal system to study replication-independent histone dynamics. Canonical H2A and all variants were deposited in the nuclei of fully grown oocytes. Despite, only histone H2A. X was abundant in the pronuclei of one-cell embryos after fertilization, in contrast with the low abundance of histone H2A and the absence of H2A.Z [87]. Fusion protein experiments using H2A, H2A.Z, and macroH2A fused with the C-terminal 23 amino acids of H2A.X showed that the C-terminal amino acids of H2A.X function specifically to target this variant histone into chromatin after fertilization, and that the absence of H2A.Z and macroH2A from the chromatin is required for normal development. These results suggest that global changes in the composition of histone H2A variants in chromatin during oocyte maturation play a role in genome remodeling after fertilization. Microinjection of mRNA for Flag-tagged histone H3.3, but not Flag-tagged canonical histones H3.1 and H3.2, led to incorporation of the histone into the chromatin of growing and fully grown oocytes. Oocyte-specific deletion of histone cell cycle regulator (HIRA), a histone chaperone required for H3.3 incorporation, abolished incorporation of microinjected H3.3 and led to chromatin decondensation accompanied by signs of DNA damage [88]. HIRA is necessary for normal gene expression and de novo DNA methylation during oocyte development.

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These studies provide further evidence that histone replacement in postmitotic cells is physiologically important. Such exchange could be important for cell-typespecific transcription and plasticity in response to external cues as well as for maintaining genome integrity. It will be interesting to determine whether the modifications of H3.3 at other residues also influence the exchange of this histone variant, and whether the exchange of other histones, such as H2A, is also regulated by modifications such as methylation and ubiquitination.

Histone H3 Lysine-4 Trimethylation Histone H3 lysine-4 trimethylation (H3K4me3) is a type of histone modifications that associates with eukaryotic gene promoters and poises them for development- or environment-triggered transcriptional activation. The H3K4me3 level was low in growing NSN oocytes (Fig. 9C and E), but was significantly increased in fully grown SN oocytes (Fig. 9D and F), suggesting that H3K4me3 accumulation is an important aspect of oocyte epigenetic maturation. After ovulation, H3K4me3 localized to the condensed chromosomes in metaphase II (MII)-arrested oocytes, and was inherited by the female pronucleus after fertilization. In mammalian cells, H3K4 trimethylation is mediated mainly by the SETD1 complex and mixed-lineage leukemia (MLL) family of proteins [89]. CXXC finger protein-1 (CFP1), encoded by the Cxxc1 gene in mice, is a key component of the eukaryotic SETD1 complex [90,91]. CFP1 binds with DNA using its CXXC finger domain and recruits SETD1 to specific genome regions [92]. In mammalian oocytes, trimethylation of histone H3 at lysine-4 is mainly mediated by SETD1-CFP1 and MLL2. The H3K4me3 level was remarkably decreased, but not completely abolished, in MLL2- or CFP1-deleted oocytes. CFP1 deletion affected the expression of a wide range of maternal genes, disregarding their original expression abundances. Similar to CFP1 deletion, MLL2 deletion in oocytes led to female infertility [93]. In contrast to H3K4me3, the H3K4me1 and H3K4me2 levels were not affected by CFP1 deletion, but were downregulated in MLL2-deleted oocytes [93]. Therefore, MLL2 may primarily be responsible for maternal H3K4me2 accumulation and indirectly affects H3K4me3 in oocytes, whereas the CFP1-SETD1 complex is the major methyltransferase that directly generates H3K4me3 during oogenesis. A balanced H3K4 methylation status is essential not only for maintaining transcription in the growing oocyte, but also for triggering zygotic genome activation (ZGA) (Fig. 10). Histone H3K4me3 is enriched in the female pronucleus after fertilization and is only start to be detected in the male pronucleus at the late zygotic stage [94]. In developing oocytes and zygotes, MLL2, MLL3, and

MLL4 were being reported to contribute to H3K4 methylation [93,95], whereas KDM1A (lysine demethylase), KDM1B, and KDM5B were reported to demethylate H3K4 [96,97]. Deletion of these KDMs caused defects in oogenesis and ZGA [98–101]. H3K4me3 deposits gene promoters as broad peaks on oocyte genome, but is promptly removed by KDMs after fertilization. As a result, only narrow and sharp H3K4me3 peaks remained on the genome of two- to four-cell embryos. This largescale H3K4me3 removal from maternal genome is necessary for successful ZGA. Although the removal of bulk H3K4me3 from zygotic chromatin is a key step that leads to ZGA, the proper deposition of H3K4me3 at the promoters of certain important zygotic genes (as narrow peaks) are also required. Despite, results of this study and other reports indicated that insufficient maternal H3K4me3 accumulation also caused failure of preimplantation embryonic development: maternal Mll2 knockout embryos arrest at the two-cell stage [93]; inhibition of histone H3K4 methytransferase activity by overexpressing a dominant negative histone H3 mutant (K4-to-M mutation) in oocytes also blocked ZGA and impaired embryonic development [95]. Collectively, these results suggest that the balanced H3K4 modification mediated by both CFP1 and KDMs is crucial for the acquisition of oocyte competence.

DNA Methylation and Demethylation in Oocytes During follicle growth, the oocyte genome is methylated and acquires the maternal imprint. CFP1-mediated H3K4 methylation was reported to be important for maintaining DNA methylation levels in oocytes. Global DNA methylation level was decreased in the CFP1deleted oocytes. Particularly, the methylation of maternally imprinted genes declined in the CFP1-deleted oocytes. Notably, the DNA methyltransferases (DNMTs) required for DNA methylation, including Dnmt1, Dnmt3a/b, and Dnmt3l, were all downregulated in the CFP1-deleted oocytes. These results indicate that CFP1 deletion in oocytes impairs the expression of DNMTs, and causes sequence-nonselective defects of de novo DNA methylation. Maternal TET3 converts the 5mC to 50 -hydroxymethylcytosine (5hmC) for DNA demethylation after fertilization, and is crucial for the developmental competence of embryos. In the CFP1deleted zygotes, Tet3 mRNA was downregulated, and the 5mC to 5hmC transition was blocked, particularly in the male pronucleus. These results demonstrate that by affecting H3K4 trimethylation, CFP1 regulates the transcription of other maternal epigenetic regulators during oocyte growth, and renders developmental competence to fully grown oocytes.

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CPL components

H3K4me3

SETD1

DNMTs

Transcription

CFP1

Exchange

BTG4

H3K4me3

SETD1

TET3

Transcription activation

CFP1

Deposition

5mC

5hmC

Histone variants

H3K4me3

mRNAs

FIG. 10 Summary of CFP1 function related to epigenetic regulation in oocyte development and zygotic genome activation. By facilitating histone H3K4 trimethylation and histone exchange in oocytes, CFP1 maintains the expression of key oocyte genes, including those encoding for DNA methyltransferases (DNMTs), cytoplasmic lattice (CPL) components, MZT licensing factor BTG4, and DNA demethylase TET3. These CFP1 regulated factors are involved in the establishment of maternal genome imprinting and cytoplasmic organelle distribution during oocyte maturation, as well as maternal transcript clearance and DNA demethylation after fertilization. Furthermore, maternal CFP1 ensure prompt histone deposition and zygotic genome activation in 1–2-cell embryos.

In addition to accumulation of maternal DNA imprinting, the balanced DNA methylation and demethylation is equally important for oocyte epigenetic maturation. Recent studies indicated that CRL4DCAF1-mediated TET activation is crucial for the acquisition of oocyte competence [102]. CRL4 bound to the TET catalytic domain through its substrate adaptor DCAF1 and activated TET by monoubiquitination at a conserved lysine site. By regulating TET-mediated 5hmC generation in oocytes, CRL4DCAF1 maintained oocyte genome DNA methylation levels and regulated the expression of essential oocyte genes, including Nobox, Figla, Sohlh1/2, and Lhx8, in both dormant oocytes of primordial follicles and activated oocytes of growing follicles [102]. By activating TET3 in the male pronucleus, CRL4DCAF1 also acted as a maternally derived zygote reprograming factor after fertilization. Oocyte-specific Ddb1 or Dcaf1 knockout female mice were infertile, because their oocytes contained hyper-methylated genomic DNAs and failed to achieve epigenetic maturation. A following up study provided evidence that CRL4DCAF1 was essential for maintaining oocyte survival and DNA methylation balances, not only those in dormancy at the primordial follicle stage, but also in awakened oocytes within growing follicles [103]. Interestingly, the oocyte-specific Ddb1 or Dcaf1 knockout

mice had ovulation defects even before oocyte exhaustion. CRL4DCAF1 within oocytes was required for cumulus expansion and ovulation-related somatic gene expression in a cell nonautonomous manner. The GCs that surrounded these Ddb1 or Dcaf1-deleted oocytes exhibited increased rates of apoptosis and showed poor responses to ovulation signals. The underlying mechanism was that Ddb1 or Dcaf1 deletion impaired the expression of genes encoding for oocyte-derived paracrine factors (Gdf9, Bmp15, and Fgf8) due to hypermethylation of their promoter regions [102]. Indeed, in oocytes that were isolated from Ddb1fl/fl; Gdf9-Cre mice, Gdf9, Bmp15, and Fgf8 mRNA levels were all decreased. These results suggested that epigenetic status in oocytes also regulated the GC functions in a cell nonautonomous manner.

Polycomb-Repressive Complexes Polycomb group proteins are evolutionarily conserved transcriptional repressors that were originally identified in Drosophila as factors required for the maintenance of transcriptional silencing of selected genes during embryonic development. Mammalian polycomb proteins function in two major complexes, polycomb-repressive complex 1 (PRC1) and PRC2, that catalyze

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monoubiquitination of H2A (H2A119ub1) and trimethylation of H3K27 (H3K27me3), respectively [1]. It has been shown that methylation by the PRC2 component enhancer of zeste homolog 2 (EZH2) and recognition of H3K27me3 by EED are required for propagation of the repressed state. H3K27me3 further recruits canonical PRC1 complexes containing different CBX and PCGF proteins. Finally, PRC1 complexes repress transcription by compacting chromatin and/or blocking RNA polymerase elongation through H2A119ub. Various PRC components are expressed in oocytes and maternally provided to the embryo. RING1 and RNF2, catalytic subunits of PRC1, serve redundant functions during oocyte maturation and are essential for proper ZGA. Genetic ablation of both paralogs results in massive transcriptional misregulation during oocyte growth, and developmental arrest at the two-cell stage of embryogenesis. Exchange of chromosomes between control and Ring1/Rnf2-deficient metaphase II oocytes reveal both cytoplasmic and chromosome-based contributions by PRC1 to embryonic development. Therefore, PRC1 acts in oocytes to establish developmental competence for the following generation by silencing differentiationinducing genes and defining appropriate chromatin states. EZH2, a core member of PRC2, is directly involved in the trimethylation of lysine-27 on histone H3. In the early mouse embryo development, EZH2 was detected as a maternally inherited protein in the oocytes. EZH2 was increased with the oocyte progression from GVBD to MII, and was concentrated on the chromosomes [104]. Depletion of EZH2 led to chromosome misalignment and abnormal spindle assembly. Furthermore, ectopic expression of EZH2 led to oocyte meiotic maturation arrested at the MI stage followed by chromosome misalignment and aneuploidy. Mechanistically, EZH2 directly interacted with and stabilized BubR1, an effect driving EZH2 into the concert of meiosis regulation.

CONCLUSION The processes comprising oocyte maturation are essential for the transition from a gamete to an embryo competent to give rise to a healthy new individual. Concurrently these same processes also represent the final stage in the generation of a highly differentiated cell, the oocyte. Although this chapter’s descriptions of the processes of oocyte maturation may seem complete to the novice, those working this field will recognize that the story is incomplete—a work in progress—with many key questions remaining to be resolved. What are the mechanisms of meiotic resumption-coupled mRNA translation and degradation? Although several key molecules governing the starts and stops of mRNA

translation in oocytes are known, major questions regarding their activation and function remain. What are their molecular partners and who determined their selective regulation of maternal mRNAs? How does their distribution and organization in the 30 -UTR affect mRNA stability? In addition, the identities of putative epigenetic regulators promoting histone exchanges and modifications in oocytes or signaling the genome remodeling after fertilization are still unknown. Disrupting histone modifications directly affected chromosome alignment and separation, as well as meiotic cell cycle progression, but the underlying mechanisms are poorly investigated. Vital maternal effect genes and their biochemical functions in regulating MZT remain to be discovered. Answers to these, and many other related questions are essential for understanding the fundamental mechanisms of reproduction and thus critical for devising novel approaches to fertility control and reducing the incidence of birth defects, as well as facilitating production and use of totipotent stem cells, efficient domestic animal production, and propagation of endangered species.

Acknowledgments H.Y.F. is funded by the National Key Research and Development Program of China (2017YFSF1001500, 2016YFC1000600) and National Natural Science Foundation of China (31528016, 31371449, 31671558).

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C H A P T E R

13 Gene Expression During Oogenesis and Oocyte Development Mo Li, Jie Yan, Xu Zhi, Yun Wang, Jing Hang, Jie Qiao INTRODUCTION An oocyte, as one of the largest cells in the body, is a female gametocyte for reproduction. It undergoes dynamic processes involving complicated regulatory mechanisms with various stages during the generation of oocyte and subsequent maturation and development. The female primordial germ cell (PGC) goes through mitosis, forming oogonia in the fetal ovaries. During the processes of oogenesis, the oogonium divides and enters meiosis I stage, becoming the primary oocyte, which ceases at the prophase stage of meiosis I. Upon puberty, the primary oocyte finishes meiosis I, and divides into a haploid secondary oocyte and an extruded first polar body. Only a few primary oocytes are recruited, and only one matures that ovulated during a menstrual cycle. When this secondary oocyte enters meiosis II, it is arrested at the metaphase of meiosis II (MII) stage until fertilization. During this time, the process of meiosis is terminated and the second polar body is extruded [1]. The whole process of folliculogenesis and ovulation is controlled by follicle-stimulating hormone (FSH) and luteinizing hormone (LH), which are secreted from the pituitary gland and controlled by estradiol, progesterone, inhibin, and others, mostly produced by the ovary [2]. Therefore, communication between the ovaries and the hypothalamic-pituitary unit is essential for periodic ovulation. The gene expression patterns in oocytes presenting in follicles at different development stages are crucial for investigating biological mechanisms controlling mammalian oogenesis and oocyte development. This chapter focuses on gene expression and regulation during oogenesis and oocyte maturation, which will be critical for optimal fertilization and embryo development. It includes the transcriptome and proteome profiles and their modifications, as well as the influencing factors during different

The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00013-3

stages [3–7]. We also summarize the epigenetic hallmarks of PGC and oocyte development to comprehend the epigenetic modification patterns [8–10]. As the dysregulation of gene expression will lead to pathological outcomes, which are directly correlated with defective chromosome condensation and segregation, delayed maturation progression, granulosa cell tumor, and others in female reproduction [11,12], we include this clinical relevance in our chapter. This data set of the chapter may promote an improved understanding of the reproduction essence, and benefit the design of optimized ovarian stimulation protocols, and thus the progression of reproductive medicine.

GENE EXPRESSION DURING OOGENESIS Molecular Regulation of Oogenesis Gene expression and regulation during oogenesis contribute to material basis for female gametes and subsequent fertilization. This presents the developmental nature of germ cells and lives. As a prolonged developmental process, oogenesis is cautiously regulated, leading to the production of functional oocytes. Oogenesis in mammals mainly encompasses three crucial periods: proliferation and migration of PGCs, formation of primordial follicles, and activation and growth of follicles [13]. These events lead to the production of developmentally competent oocytes capable of fertilization in the future. The differentiation and development of PGCs are characterized by dynamic and remarkable alteration in gene expression, most of which is regulated by a series of transcriptional factors. Using laser microdissection, Markholt et al. isolated distinct populations of oocytes from early follicles in the ovary [14]. Their work shows more than 6000 genes

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that are significantly expressed. These genes are classified into seven categories, such as “RNA binding,” “translation initiation,” and “structural molecule activity,” which highlight the complex of gene network during oogenesis. The bidirectional interactions between germ cells and the surrounding granulosa cells (GCs) are also necessary to form an ovarian follicle. Studies on gene expression and regulation are crucial to develop a better understanding of mechanisms of follicles maintenance and activation, and thus facilitate reproductive biology.

Gene Expression During PGCs’ Migration and Proliferation Human PGCs are precursors of oocytes and are crucial for maintenance of the species. In female gonads, PGCs differentiate into oogonia by mitosis [15,16], which undergo massive proliferation by mitotic divisions from 6 to 8 weeks gestation [17] and then become primary oocytes that arrest at the prophase of the first meiotic division. During this phase, PGCs migrate to the gonadal ridges from the hindgut by crossing the dorsal mesentery. A series of factors, including the transforming growth factor beta (TGFβ) family members and germ cell-derived transcription factors [18,19], are involved in this process. The expression levels of BMP2 and BMP4 increase in mice PGCs, while the expression level of Activin increases in human PGCs. The transcriptional factors that play important roles in survival, migration, and proliferation of PGCs include SOX17, BLIMP1, PRDM14, OCT4, NANOG, FIGα, NANOS3, and DND1 [20]. Through incomplete division of the cytoplasm (incomplete cytokinesis) during mitosis, oogonia form germ cell cysts that will develop into ovarian follicles [21]. The growth phase of oogenesis is initiated when oogonia finish their mitotic division, together with their enlargement to form primary oocytes and meiosis initiation. During 11–20 weeks of embryonic life before birth, the first meiotic division in primary oocytes is initiated in humans. During this process, several RNA binding proteins, exemplified by DAZL and BOLL, are involved in different stages of human meiotic division [22]. PGCs react upon signals secreted by neighboring somatic cells through receptor/ligand interactions to facilitate their migration through the hindgut and into the developing gonads. Released by granulosa cells, KITL recognizes its receptor KIT on the oocyte, inducing gonad development. Using single-cell RNA-sequencing techniques (RNA-seq), a recent study revealed that several signaling pathways are coordinately and reciprocally enriched between PGCs and their gonadal niche cells [23]. For example, the NOTCH signaling pathway is specifically activated in gonadal somatic cells, while the BMP

signaling pathway is activated in PGCs by BMP2 secreted from the neighboring granulosa cells [23].

Gene Expression During Primordial Follicle Formation After the PGC migration and proliferation, the oocyte enters the diplotene stage during the prophase of the first meiosis and is arrested in a prolonged resting phase [21]. In humans, primordial follicle formation occurs during the second trimester of fetal development and is completed before birth [16]. During primordial follicle formation, germ cell cysts are interrupted and the primary oocyte is surrounded by ovarian stromal (pregranulosa) cells to form the single-layer primordial follicles. The STRA8 (stimulated by retinoic acid gene 8) signaling, dependent on the stimulation retinoic acid signaling, is involved in the initiation of meiosis in both human and mouse ovaries [21]. A study in model organism demonstrates that estrogen and the corresponding receptors are involved in the process of follicle cyst breakdown and primordial follicle formation, as well as TGFβ superfamily members, particularly GDF9 and BMP15 [24]. However, factors and signaling pathways in humans remain to be explored. The formation of the primordial follicles is coupled with the apoptosis of a large number of oocytes, since different pro- and antiapoptotic proteins have been implicated as significant in germ cell fate determination during primordial follicle formation [25]. These proteins include B cell lymphoma-leukemia (BCL) protein family members BCL2 and BCLX and proapoptotic protein BAX. It has been demonstrated that an absence of BCL2 or BCLX results in a reduced number of primordial follicles, whereas the loss of BAX or Caspase 2 activity leads to an increase in the primordial follicle pool [19]. Primordial follicles presented at birth indicate the reservoir of germ cells available during female reproductive life. Thus, mutation or dysfunction of any of the genes involved in PGCs’ migration and survival, regulation of follicle formation, and apoptosis can lead to the diminution of the germ cell pool and in turn affect a woman’s reproductive span.

Gene Expression During Follicle Activation and Development The signaling interaction between oocytes and surrounding granulosa cells guarantees the highly complex and dynamic follicle activation and development. Before activation, the primordial follicles remain dormant until puberty and then undergo activation as cohorts of continuously recruited follicles. After primordial follicle activation, the oocytes enlarge, and flattened granulosa cells develop into a cuboidal shape and go into a proliferative state. Activation of the primordial follicles is regulated by

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a series of genes productions. PTEN/PI3K signaling has been identified as crucial for transition from primordial to primary follicles. Phosphatase and tensin homolog deleted on chromosome 10 (PTEN) leads to inactivation of phosphatidylinositol 3 kinase (PI3K), a serinethreonine protein kinase that stimulates cell proliferation and suppresses the expression of pro-apoptosis factor forkhead box O3 (FOXO3). Studies on mouse and human ovaries demonstrate that manipulation of the PTEN/ PI3K pathway by inhibition of PTEN and activation of PI3K induces primordial follicle activation [26]. Another signaling pathway is mediated by the Tsc/mTORC1 (tuberous clerosus complex 1 and mammalian target of rapamycin complex 1) signaling transduction [27]. Despite the derivation from animal studies, many other vital factors have been proved as well, including KIT ligand and its receptor, neurotrophins, transcriptional factors Nobox and Sohln1, and the members of TGFβ family BMP4, BMP7, and anti-Mullerian hormone (AMH) [19]. As follicles grow, the zona pellucida forms outside primary oocytes and cuboidal somatic cells gain additional layers, designating the progression to secondary and antral stages. Only a few oocytes progress through maturation in every menstrual cycle. The involved factors include GDF9 and BMP15 from the TGFβ family, and TATA-binding protein 2 (TBP2) and TATA box-binding protein (TBP)-associated factor (TAF4B) [28]. The early developmental stage of ovarian follicles is follicle stimulation hormone (FSH)-independent and controlled by locally secreted factors, and then transfers to an FSHdependent manner when reaching the antral stage. Just before ovulation, the primary oocyte completes the first meiotic division, resulting in the formation of the secondary oocyte and the extrusion of the first polar body, a small non-functional cell that subsequently degenerates. The second meiotic division begins immediately

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at ovulation and completes upon fertilization, leading to the creation of a haploid ootid with the extrusion of the second polar body [1]. In addition to the genes expressed in the oocyte, transcriptional regulators expressed in the surrounding cells have been shown to play important roles during folliculogenesis. Oocyte-granulosa bidirectional communication is one of the crucial mechanisms of oocyte acquisition of developmental competence. Among the factors that orchestrate oocyte development, activation of NOTCH signaling in granulosa cells (GCs) by ligands from adjacent germ cells [29] highlights the role of NOTCH signaling in oocyte-controlled proliferation and differentiation of GCs. Moreover, members of the TGF-β superfamily, the oocyte-derived factors GDF9 and BMP15, have been shown to be involved in oocyte maturation, GC proliferation, and cholesterol biosynthesis [30]. In addition, GC-derived signaling, KITLG and its receptor KIT, previously implicated in paracrine signaling, is expressed in GCs and oocytes, respectively, and plays a crucial role in primordial follicle activation [31]. Over the past decades, substantial progress has been made in elucidating the molecular mechanisms governing the gene expression of oogenesis. Due to the inaccessibility of human ovaries, previous knowledge was obtained mainly based on studies in mice. The crucial roles of proteins expressed during oogenesis have been elucidated through knockout approaches or targeted deletions in mice. Nevertheless, many questions remain to be answered. Accumulating evidence has shown that human PGCs have a unique gene expression network different from that of mice. Technique advances, represented by such as single-cell resolution analyses, are expected to contribute to the collaborative effort to strengthen our understanding of oogenesis in humans (Fig. 1).

FIG. 1 Molecular mechanisms of oogenesis. Oogenesis starts with PGCs proliferation and mitosis, followed by oogonia meiosis and germ cell meiotic arrest. Subsequently, primordial follicles are activated to form primary follicles and developed into growing follicles. This process is accurately regulated by a set of molecules at certain times. II. OOCYTE MATURATION AND OVULATION

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GENE EXPRESSION PROFILE DURING OOCYTE MATURATION The mammalian ovary contains thousands of primordial follicles, which form during fetal development or shortly after birth. Each primordial follicle contains a single oocyte, which is
10–15 μm in diameter (in mice) and arrested in diplotene of the first meiotic prophase. Oocytes are derived from primordial follicles. After birth, mammalian oocytes are arrested at the diakinesis of prophase I in the ovary, which is known as the germinal vesicle (GV) stage. Meiosis resumes when the germinal vesicle breaks down (GVBD) after stimulation by luteinizing hormone (LH) at the beginning of puberty [32]. Resumed meiosis I progresses through metaphase I (MI) when homologous chromosome pairs move together along the metaphase plate, followed by segregation of the homologous chromosomes. After the exclusion of the first polar body, meiosis II initiates without DNA replication. This process is similar to mitosis and involves equational segregation of sister chromatids after cohesin degradation. As a result, the number of DNA copies halve, while the number of chromosomes remains invariant [33]. To further understand the molecular basis of oocyte maturation, the identification of specifically expressed during oocyte maturation, has been performed extensively. In recent years, the development of transcriptomic analysis has permitted the evaluation of global gene expression change in mouse and human oocytes [3]. In a microarray analysis, Cui XS et al. examined 12,164 genes in GV- and MII-stage oocytes in mice. Their work showed that when comparing the GV stage with the MII stage in mouse oocytes, more than 1600 genes were upregulated in GV-stage oocytes, while more than 1900 genes were upregulated in MII-stage oocytes [34]. In addition, the genes increased in the MII oocytes may take part in amino acid metabolism, G protein-coupled receptor signaling, DNA replication, and expression of signaling molecules [34]. A previous report showed that mRNAs synthesized during oocyte growth contribute to early embryonic development before zygotic genome activation. Maternal mRNAs in mouse oocytes are unusually stable during the growth phase, with an average half-life of approximately 10–14 days, compared to hours or minutes in somatic cells [35]. Su et al. clarified that this destruction of transcripts during the GV to MII transition is a selective rather than indiscriminate process. Moreover, they found out that the dramatically degraded transcripts were associated with GV-stage meiotic arrest, oxidative phosphorylation, energy production, and protein synthesis and metabolism, whereas the stable transcripts were mainly involved in signaling pathways, which were important for maintaining the MII oocyte characteristics. Elevated

concentration of cAMP, which is produced from ATP, maintains meiotic arrest in oocytes. Therefore, the transcripts associated with oxidative phosphorylation and energy production were downregulated in MII oocytes in order to acquire meiotic competence to complete meiotic division [36]. Furthermore, a similar transcriptome change was observed in human oocyte maturation. For example, the gene expression pattern of human oocytes at the GV and MI stages has been compared and the results showed that minor changes occur during these two immature stages. By contrast, MII-stage oocytes showed a marked overexpression of 444 genes and a significant downexpression of 803 genes in comparison with immature stage oocytes, suggesting that the dramatic change of gene expression pattern occurred at the last stage of oocyte maturation. Assou et al. found that plenty of representative genes that were associated with the key biological processes such as mitotic cell cycle, DNA replication, and fertilization, appeared in the MII stage. They also observed that in oocytes, the expression level of 1514 genes increased at least threefold, including Dazl, Ddx4/Vasa, and Dppa3/Stella. These genes have been considered to be expressed specifically in mammalian primordial germ cells. Consistent with previous studies, numerous well-recognized actors of meiosis were also detected—for example, the components of the maturation-promoting factor (MPF), spindle checkpoint, anaphase promoting complex (APC/C), and meiosisspecific sister chromatid arm cohesin. As expected, the genes that are considered to be specifically expressed in oocytes such as ZP1, 2, 3, and 4, Gdf9, Bmp6, Fgfr2, Hdac9, and the oocyte-specific H1 histone H1FOO were also observed. Interestingly, they also revealed that several genes that had been reported to be expressed in the male germ cells were also highly expressed in oocytes, including Aurkc, Sox30, and Spag16/PF20 [37]. In addition, the proteasome is known to interact with chromatin during multiple stages of transcription. Some studies also observed that proteasome-related genes played significant roles during human oocyte maturation. BARD1BRCA1 heterodimer has been considered as an E3 ubiquitin ligase. Gasca et al. found that the multiprotein E3 ubiquitin ligase complex containing BARD1, BRCA1, and BRCA2 was involved in human oocyte maturation [37]. These proteins play crucial roles in regulating cell cycle progression, DNA repair, and gene transcription. Although it has been discovered that the differentially expression genes had unique significance, most of the genes are poorly investigated; this indicates that these genes may participate in the maturation of oocyte cytoplasm and/or nucleus. To further identify the genes essential for oocyte development, Kocabas et al. compared human oocytes transcriptome with a mixture of RNA which contains

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10 different human tissues. Their results showed that 15,331 transcripts increased significantly and 7074 transcripts decreased significantly in the oocytes. Among the upregulated genes, the roles of 1430 genes are unknown. In these intersecting significantly upregulated genes between human and mouse oocyte transcriptomes, the authors found that a group of 1587 genes may have conserved function in mammalian oocytes. Most of these genes participated in key biological processes during oocyte maturation, including the period from the first meiotic division to MII arrest and various cell cycle controls. Recently, single-cell RNA-seq techniques had paved the way to analyze the gene expression profile of human oocytes by single-cell and single-base resolution [4]. Using these methods, Yan et al. revealed that 13,986 RefSeq genes, 7214 known lncRNAs, and 1487 novel lncRNAs were maternally expressed in human mature oocytes [5]. Meanwhile, the transcriptional activity has been reported decreased in fully grown mouse oocytes [38]. During this silent period, the oocytes depend on prestored RNAs for its maturation process, and about half of transcripts were destroyed during oocyte maturation [39]. However, which transcripts have been degraded during the transition from the GV to the MII stage was unknown. Using microarray analysis, Su et al. uncovered the destruction of transcripts in oocytes during the transition from the GV to the MII stage. These data demonstrated that it was a selective rather than a random process at these stages: transcripts that participate in the progression of oxidative phosphorylation, energy production, and protein synthesis or metabolism were dramatically degraded during oocyte maturation. Other studies showed that the global expression pattern of oocytes maturation has been identified as particularly sensitive to the maturation environment. In mammals, cell adhesion, cellular homeostasis, cell–cell interactions, and mRNA stability or translation could lead to different mRNAs expression between in vivo and in vitro oocyte maturation [40]. In rhesus monkeys, a relatively small number of transcripts was found to be expressed differently in these two cases (n ¼ 59), and most (90%) of them were upregulated in the in-vitro matured (IVM) oocytes. A similar performance was found in cattle [41]. However, in mice and humans, the transcriptomes of the in vivo-matured oocytes were detected to express a greater number of genes, most (70%) of which were upregulated in the in vivo-derived oocytes [42,43]. Furthermore, bioinformatical analysis of the upregulated genes revealed that with the expression of OCT4, the key transcription factors, these oocytes preferentially express genes associated with adverse pathways such as oxidative phosphorylation, mitochondrial dysfunction, and apoptosis [43].

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THE REGULATION OF GENE EXPRESSION DURING OOCYTE MATURATION mRNA Stability in Oocytes After a period of active transcription during growth, the nuclei of mammalian oocytes become transcriptionally inactive [44]. Transcription ceases at the onset of meiotic resumption and becomes fully activated in the zygotic genome activation (ZGA) stage, which occurs at the two-cell stage in mice and the four- to eight-cell stage in cattle and humans [5]. Oocyte mRNA is extremely stable, and the regulation mechanism of mRNA stability in the oocyte is clarified as follows: (1) poly(A) tail length and 30 terminal uridylation: It has been observed that the ratio of poly(A) to total RNA is higher in oocytes than in somatic cells [45]. Some NGS studies suggest that the regulation of poly(A) tails in oocytes shapes the translatomic landscape [46]. Another study shows that 30 terminal uridylation of mRNA mediated by TUT4 and TUT7 sculpts the mouse maternal transcriptome by eliminating transcripts during oocyte growth. In summary, poly(A) tail length and 30 terminal uridylation have essential and specific functions in shaping a functional maternal transcriptome [47]. (2) Localization: The majority of mRNAs are subcellularly localized. Dozens of detected mRNAs are limited to the spindle, nucleus, and perinuclear regions, or they are membrane-associated. In mice, the nucleus of a fully grown oocyte shows the high poly(A) RNA signal that is retained in the chromosomal area, and the signal disappears as the oocyte proceeds through meiosis to the MII stage. Tight correlations between mRNA distribution and subsequent protein localization as well as function indicate major roles for mRNA localization. (3) Ribonucleoprotein particles (RNPs): RNPs are known to regulate translation and localization of mRNA in mammalian oocytes. RNAs, together with numerous RNA-binding proteins concentrated in the nucleus, at the spindle, and in the cytoplasm, are believed to play a role in mRNA storage and metabolism [48–50]. RNP aggregates contain poly(A) RNAs and cytoplasmic polyadenylation element-binding protein 1 (CPEB1), DEAD-box helicase 6 (DDX6), and Y-box-binding protein 2 (MSY2). These aggregates disperse during meiotic maturation and release maternal mRNAs for translation [48]. (4) Subcortical Maternal Complex (SCMC): Subcortical complexes share components with P-bodies (processing bodies) including several RNA-binding proteins, such as CPEB1, DDX6, and MSY2, with their function as a storage compartment [48].

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Translation Regulation in Oocytes During oocyte maturation, numerous mRNAs will be translated into proteins, specifically required for oocyte function, while another set of mRNAs will be translated later in the embryonic period, after fertilization. In mouse oocytes, large numbers of maternal mRNAs are recruited to polysomes during the progression of the meiosis. Approximately 7600 mRNAs have been shown to be translated during oocyte maturation by microarray analysis of RNAs purified from the polysomes of oocytes at the GV, MI, and MII stages [51]. These mRNAs are classified into three categories according to the patterns of polysome recruitment in GV and MII oocytes. Around 4800 mRNAs are constitutively recruited on polysomes (class I). In contrast, a set of 1500 mRNAs recruited to polysomes decreases more than twofold during oocyte maturation (class II), and around 1300 mRNAs on polysomes increase by more than twofold during maturation (class III). Of these mRNAs, the translation of some is known to be essential for the progression of oocyte maturation and development. For instance, Cyclin B1 protein synthesized from stored mRNA forms a complex termed MPF with preexisting Cdc2 kinase, and immediately activates the Cdc2 [52,53]. MPF phosphorylates hundreds of substrates to promote meiotic cell cycles properly as well as cytoplasmic and nuclear maturation [54].

miRNA in Oocytes miRNAs play a vital role in regulating and manipulating maternal mRNAs for normal oocyte maturation. Genes essential for miRNA biosynthesis include Dicer, AGO2, and DGCR8. In addition to expressing full-length Dicer, mouse oocytes produce an oocyte-specific form of Dicer (Dicero) [55]. Dicero is particularly efficient at generating endogenous small interfering RNAs (siRNAs) and is essential for mouse oocyte function [55]. Doublestranded siRNAs are generated through the Dicer/ AGO2 pathway, but do not rely on DGCR8. Knockout of dicero causes abnormal spindle formation and chromosomal organization [56], similar to the original Dicer-zp3-cKO and AGO2-zp3-cKO mice, thus confirming these earlier studies [55–57]. However, knockout of DGCR8 does not affect the oocytes’ maturation [58]. These earlier studies have proven a requirement for siRNA but not miRNA pathways for mouse oocyte maturation. Several interesting studies have been conducted in bovine oocytes that support an important role for miRNAs during oocyte maturation. Bta-miR-155, bta-miR222, bta-miR-21, bta-let-7d, bta-let-7i, and bta-miR-190a are expressed in bovines, while P53 is targeted by the let-7 family of miRNAs [59]. Another study in bovines examined the role of miR-212 in the regulation of FIGLA.

miR-212 is highly expressed in bovine GV oocytes with a tendency to increase in the cleavage stage of the embryo up to the eight-cell stage—the time at which the bovine embryo undergoes that transition from maternal to zygotic (embryonic) gene regulation. miRNAs in granulosa cells have been found to play a significant role in regulating oocyte maturation [60]. For instance, miR-21 and three miRNA clusters (miR-183-96182, miR-132-212, and miR-424-450-542) were preferentially enriched in granulosa cells, involved in rescuing granulosa cells of the preovulatory dominant follicles from undergoing apoptosis. In mice, increased levels of miR-21, miR-132, and miR-212 in granulosa cells have been found following the LH surge [61,62]. Carboxyterminal binding protein 1 (Ctbp1), as a putative target of miR-132/212, interacts with steroidogenic factor-1 (sf-1), indicating a potential role in regulating steroidogenesis in granulosa cells. These studies show that a loss of activity associated with specific miRNAs in vivo can disturb ovulation. Moreover, miRNAs in granulosa cells have been found to transfer to oocytes across gap junction channels, indicating their potential role in the oocyte [63,64].

Oocyte Proteome The oocyte contains a full complement of maternal proteins required for maturation, fertilization, transition to zygotic transcription, and early embryogenesis. Many of these proteins in mouse oocytes have yet to be characterized. In a recent study, Wang et al. identified 2781 proteins present in GV oocytes, 2973 proteins in MII oocytes, and 2082 proteins in zygotes through semiquantitative MS analysis [65]. Furthermore, the results of the bioinformatics analysis indicated that different protein compositions are correlated with oocyte characteristics at different developmental stages. Transcription factors and chromatin remodeling factors are more abundant in MII oocytes. In specific transcription factors TCL, family proteins TCL1, TCL1b1, TCL1b2, and TCL1b3 are more prevalent in MII oocytes than in GV oocytes. Also, epigenetic modification-related proteins are expressed more abundantly in MII oocytes. During the MII stage, oocytes express four histone demethylases—Padi6, Jmjd3, Fbxl10, and Aof2; five histone acetyltransferases— Nat11, Hat1, Taf15, Btaf1, and Taf7; six histone deacetylases—Hdac2, Hdac6, Sirt2, Sirt5, Sap30, and Satb1, and eight histone methyltransferases—Men1, Mll3, Mllt10, Mllt4, Nsd1, Prmt1, Dpy30, and Carm1. In addition, more DNA repair proteins are expressed in MII oocytes than in GV oocytes, such as proteins in nucleotide-excision repair and DNA replication [66–68]. These findings suggest a sophisticated regulation of gene production in oocytes.

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However, global oocyte proteomes of human oocytes during oocyte maturation have been limited because of the limited quantity of oocytes used and the limitations of the technology. In a recent study, Virant-Klun et al. identified 2154 proteins by single-cell proteomics of human oocytes [7]. Among these targets, 158 oocyteenriched proteins including ECAT1, PIWIL3, and NLRP7 have been identified as oocyte maturation-specific proteins. More importantly, Tudor and KH domaincontaining proteins (TDRKH) have been found to be preferentially expressed in immature oocytes. In contrast, Wee2, PCNA, and DNMT1 are enriched in mature oocytes. Thus, the development of single-cell proteomics of human oocytes will no doubt promote our understanding of the nature of oocytes and translational medicine.

EPIGENETIC DYNAMICS DURING PGC AND OOCYTE DEVELOPMENT In mammals, global epigenetic reprogramming happens during the germline development [69]. A much clearer whole view of mammalian germline development has been obtained through extensive studies using the mouse model. However, due to the scarcity of materials, knowledge on human germline development is based mainly on extrapolation from mouse research, though

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it is supplemented with sporadic immunostaining studies in humans. Recently, with the development and implementation of single-cell and low input next generation sequencing, our team and other research groups are able to profile the transcriptome and epigenome of different development stages of the germline [8,9,70,71]. This paves the way for dissecting the molecular regulation of human germline development (Fig. 2).

Epigenetic Events During PGC Development In humans, germline development begins with the specification of PGCs, the precursors of sperm and oocytes, which is expected to happen at the perigastrulation embryo stage (approximately embryonic weeks 2–3) [10,69]. During gestation (weeks 3–5), the hPGCs migrate from the yolk sac wall through the hindgut to the developing gonads. In female embryos, gonadal hPGCs remain proliferative until around weeks 10, when they asynchronously initiate the meiotic prophase. DNA methylation is an important epigenetic modification, which has a strong correlation with gene repression and genome stability maintenance. DNA methylation patterns in the genome of differentiated cells are usually stable and heritable. However, dramatic methylome reprogramming occurs in preimplantation embryo development and PGCs’ development. With the help of whole

FIG. 2

DNA epigenetic dynamics during germ cell development, oocyte growth and maturation. The higher panel shows the major events during the whole development. At female embryonic weeks 2–3, germline development begins with the specification of hPGCs, and the hPGCs migrate to developing gonads during gestation in weeks 3–5. Before weeks 4–7, the first wave of genome-wide DNA demethylation in hPGCs takes place; it reaches the lowest point at week 10 and is maintained through the 17 weeks of PGCs (blue full line). The DNA demethylation dynamics in mice during E10.5–13.5 are comparable to humans during developmental weeks 5–17. By birth, female PGCs have become primary oocytes, and arrest at the meiotic prophase I. In mice, de novo methylation is initiated in oocytes at around P10, occurring in parallel with the follicular growth phase of oogenesis; this is largely completed by the GV stage, at approximately P21 (orange full line). The green dotted line shows the hypothetical pattern of de novo methylation in human oocyte maturation. The grey rectangles summarize chromatin reorganization and histone modification during oogenesis and oocyte maturation. The yellow rectangles indicate the main regulatory factors of DNA methylation.

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genome bisulfite sequencing (WGBS) and reduced representation bisulfite sequencing (RRBS), the dynamic genome-wide DNA demethylation pattern during PGCs’ development has been revealed at single base resolution recently [8,10,70,71]. In humans, the first wave of genome-wide DNA demethylation in hPGCs usually takes place human embryonic weeks 4–7, as at this time point, the migratory hPGCs in hindgut have already exhibited a lower level of DNA methylation compared to adjacent somatic cells. Approximately 10 weeks after gestation, the global DNA methylation levels of female hPGCs would reach the lowest point, at which the entire genome is only 6%–7% (median level) residually methylated. To our knowledge, this is the lowest level of DNA methylation that occurs in human genome regardless of cell types. However, after just one more week (i.e., at week 11), DNA methylation increased to 9% in female PGCs and maintained until week 17, suggesting that the global re-establishment of DNA methylation in female PGCs starts at 11 weeks of gestation [8]. Moreover, it has been assured through systematic comparison that the DNA demethylation dynamics in mice during embryonic day (E) 10.5 to 13.5 are comparable to those in human PGCs during developmental weeks 5–19. This implies that the epigenetic reprogramming of the germline is a fundamental and highly conserved process in mammals. In the current literature, the passive dilution mechanism is the most acceptable explanation for genome-wide DNA methylation erasure in PGC recruitment of DNMT1 to the hemi-methylated DNA during replication) and the de novo methylation enzymes (DNMT3A and DNMT3B, the major enzymes with de novo methylation activities) are repressed in both human and mouse PGCs after specification; this enables replicationcoupled DNA demethylation when PGCs proliferate [72]. In addition, recent evidence suggests that active demethylation through TET1 and TET2, which converts 5-methycytosine (5mC) to 5-hydroxymethylcytosine (5hmC), also partly contributes to global DNA demethylation, especially for imprinting regions in early PGCs development [73]. DNA methylation is important for repression of gene expression, retrotransposons, and maintenance of genome stability. The global clearance of DNA methylation observed in hPGCs includes a great loss of DNA methylation at CpG islands, transcription start sites, gene bodies, intergenic regions, repeats, and even in most imprinting regions. The loss of CpG methylation, globally uncoupled with the changes in gene expression or retrotransposon activation in hPGCs, suggests that the key purpose of demethylation is to clear the parental epigenetic memory. However, a small subset of genes that is involved with the late germ cell development and genome defense is upregulated and correlated with the removal of DNA methylation at the promoters of genes,

indicating an instructive role of DNA demethylation in PGCs’ development. DNA demethylation in both human and mouse PGCs is also accompanied by global chromatin modification reorganization. Apart from the global loss of repressive chromatin marker H3K9me2, repressive H3K27me3 is enriched during the course of mPGC development, and H3K9me3 will be retained predominantly at pericentric heterochromatin. In addition to modulating gene expression, these markers have also been implicated in the repression of retrotransposons in mPGCs [70]. Furthermore, global reorganization of repressive histone modifications is also observed in hPGCs, albeit with slightly different dynamics. Thus, it is likely that one of the purposes of germline chromatin reorganization is to safeguard genome integrity while PGCs undergo DNA demethylation to exceptionally low levels. Despite of global DNA demethylation, evolutionarily young and potentially hazardous retrotransposons, such as IAP (Intracistemal A Partical) in mice and SVA (SINE-VNTR-Alu) in humans, remain relatively highly methylated, which may contribute to their repression [8]. Notably, there are also some repeat-poor regions that can escape from global demethylation, preferably those located at enhancers, CGI, promoters, and gene bodies, which have potential abilities to represent the hotspots of transgenerational epigenetic inheritance. In human and mouse fetal germ cells (FGCs), nucleosome-depleted regions (NDRs) are found to cover a large set of germlinespecific and highly dynamic regulatory genomic elements. Notably, distal NDRs are specifically enriched for binding motifs of the pluripotency and germ cell master regulators, which indicates there is a delicate regulatory balance between pluripotency-related genes and germ cell-specific genes during fetal germ cell development [74].

Epigenetic Events During Oocyte Growth and Maturation After global DNA demethylation in early PGCs, female germlines maintain genomic hypomethylation, and the meiotic arrests through the rest of embryonic development [75,76]. By birth, female PGCs have become primary oocytes, and many oocytes undergo apoptosis at this time. In mice, the postnatal day 5 (P5) oocytes start to enlarge their cytoplasmic volume and accumulate the oocyte-specific transcription units. During this period, growth of oocytes is asynchronous, with some oocytes growing faster than others. De novo methylation is initiated in oocytes at around P10, or when the oocyte reaches at least 40–45 μm in diameter. In oocytes, de novo DNA methylation occurs in parallel with the follicular growth

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phase of oogenesis and is largely complete by the GV stage, which is at approximately P21 [77]. During the process of human oocyte maturation, CpG methylation in most part of the genome is stable, yet non-CpG methylation is dynamic and continues to accumulate [78]. In the growing oocytes from mice, DNA methylation of the imprint gene is established in an oocyte size-dependent manner, not due to aging [79]. Unlike any other types of mammalian cells, the oocyte methylome is predominantly heritable, with almost no methylation in intergenic regions [80]. Genetic studies have revealed that de novo DNA methylation occurring in oocytes is primarily through the DNMT3A/DNMT3L complex [75]. Strikingly, in oocytes, about 90% of hypermethylated regions are correlated with active transcription and, similarly, approximately 90% of hypomethylated regions are confirmed as untranscribed, demonstrating an excellent correlation between transcription and DNA methylation [81]. The most rational explanation of such strong correlation between transcription and DNA methylation is that it might be the consequence of how targeting of the DNMT3A/DNMT3L complex is regulated by histone posttranslational modifications, as growing evidence has shown that H3K4 methylation inhibits the interaction of DNMT3A/DNMT3L with nucleosomes, and H3K36me3 enhances DNMT3A activity. Moreover, it is well-known that H3K4me2 and H3K4me3 are classic marks of active promoters and CGIs, and H3K36me3 is an active transcriptional mark enriched in gene bodies, therefore building the link between transcription and de novo DNA methylation. The functional role of DNA methylation in the oocyte is not very informative. Loss of DNA methylation in the oocyte from Dnmt3a conditional mutant females has no effects on its growth, maturation, or fertilization. However, embryos conceived from those oocytes died by E10.5, largely because of the deficiency of methylation at maternal imprinted regions [82]. Dnmt3L is shown to work together with Dnmt3 family methyltransferases for de novo methylation of maternally imprinted genes in oocytes [83]. In addition, methylation at some loci contributes to gene regulation. More recently, maternal methylation has been shown to be required for differentiation and physiological function during trophoblast development [84]. The breakdown of the germinal vesicle signifies oocyte commitment to maturation. The maturation period goes from germinal vesicle to metaphase II. Histone modification, such as methylation and acetylation by specific enzymes, is a key epigenetic mechanism that controls gene expression during embryonic development. During oocyte maturation, dynamic and differential histone modification patterns have been observed, indicating a functional requirement in regulating gene activity. Acetylation of H3K9 (histone H3, lysine 9), H3K18, H4K12,

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and H4K5, and methylation such as H3k4me2, H3K4me3, and H3K9me2 increase steadily during oocyte growth. With the oocyte meiotic maturation, however, chromatin histones undergo widespread deacetylation [85]. Disruption of histone modification can lead to defective chromosome condensation and segregation, delayed maturation progression, and even oocyte aging [86].

ABNORMAL EXPRESSION OF GENES IN REPRODUCTIVE DISEASES Clinically, it was found that birth defects and lower implantation rates in humans increased with advanced maternal age, and aneuploidy in oocytes was considered to be a leading genetic reason. One hypothesis to explain this is cohesion deterioration with advanced age-related aneuploidy. The mechanism has been studied in both mouse and human oocytes [11,87–89]. The cohesion is established in premeiotic S phase during fetal development by cohesin. There was no more replenishment in the whole female fertility. The function of cohesin is to hold sister chromatids together. This is the case until puberty, when meiosis resumes and chromosomes segregate; the sister chromatid cohesion is released at anaphase I and II. For a correct segregation process, the cohesion’s cleavage time is vital. This process is regulated by a series of genes and their transcription and expression, and other factors. However, in aged oocytes, due to the premature loss of the Sgo2 protein, increased sensibility of separation, abnormal activation of SAC and APC/CCdc20, or oxidative damage, cohesion deteriorates. At transcriptional level, aging causes a reduction in Mad2 and Bub1 in human and mouse oocytes [43,90]. In the aged mice model, it was found that the level of Mad2 on kinetochores was dominantly reduced at 3 h after the resumption of the first meiosis [91]. Furthermore, in aged human oocytes, the kinetochore localization of both Bub1 and BubR1 proteins decreases accordingly [92]. Another research proved that Smc1beta also placed an important role in activating SAC during oocyte meiosis [93]. Thus, it is supposed that decreased kinetochore localization of SAC is correlated to cohesion loss with maternal age. It is probable that SAC ability is reduced in aged oocytes, which could further lead to APC/CCdc20 activation to degrade cyclin B1 and securin. These could cause sister chromatid cohesion loss [89,94]. Ultimately, the factors mentioned here, either alone or together, may influence the strength of cohesion with advanced maternal age. In addition to the influence in oocyte quality, abnormal expression and mutation of genes may induce ovary tumors. One dominant case are ovary granulosa cell tumors. These tumors are relatively rare, and account

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for approximately 3% of all ovarian malignancies. Several studies have focused on somatic mutations of different types of ovary granulosa cell tumors. In the case of juvenile granulosa cell tumors, a somatic mutation of an oncogene, gsp, is involved in approximately 30% of the tumors. Furthermore, an excessive activation AKT variation is found in a further 60% [12]. In the case of adult granulosa cell tumors, Shah et al. have reported that over 97% of cases possess a specific FOXL2 C134W missense mutation via transcriptome analysis using massive parallel sequencing or “RNA-seq,” and molecular mechanisms relating to different stages, behavior, and prognosis need to be explored [95–97]. Additionally, in the epigenetic aspect research, recent data have indicated that hypermethylation of promoters of CDH13, DKK3, and FOXL2 genes, and overexpression of EZH2 protein, are involved in the development of granulosa cell tumors. This has been proved in 30 granulosa cell tumor tissues and 30 healthy control tissues using methylation-specific polymerase chain reaction analysis [98]. More research is being developed to explore the specific genomic modification and gene expression pattern.

CONCLUSION Gene expression during oogenesis and oocyte development reflects the developmental nature of female germ cells and fundamental knowledge of our life. Gene production in oocytes contributes to the major maternal materials that are essential for fertilization, remodeling of zygotic genome, and early embryonic development. The regulations of transcription and translation described in this chapter thus provide a comprehensive understanding of these events in reproductive and developmental biology. Due to the progress of modern technology in life science, transcriptome, proteome, and methylome in oocytes have been substantially developed. Combined with molecular biology, this data set could uncover many more details for the global map of gene expression and crosstalk of key factors in female germ cells. These advances would provide clues for the optimization of obstetrics and gynecology.

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14 Ovulation: The Coordination of Intrafollicular Networks to Ensure Oocyte Release Darryl L. Russell, Rebecca L. Robker Abbreviations AREG BMP15 BTC cAMP CC CL COC COX-2 CREB DNA eCG ECM EGF EREG ERK FSH FSHR GC GDF9 hCG LH LHR MGC MMP MII NSAID OSF P4 PCOS PG PGR PKA PRKO SPRM TGFβ

amphiregulin bone morphogenetic protein 15 betacellulin cyclic adenosine monophosphate cumulus cells corpus luteum cumulus oocyte complex cyclooxygenase-2 cyclic AMP regulatory element binding protein deoxyribonucleic acid equine chorionic gonadotrophin extracellular matrix epidermal growth factor epiregulin extracellular regulated kinase follicle-stimulating hormone follicle-stimulating hormone receptor granulosa cells growth and differentiation factor 9 human chorionic gonadotrophin luteinizing hormone luteinizing hormone receptor mural granulosa cells matrix metalloproteinase metaphase II nonsteroidal antiinflammatory oocyte secreted factor progesterone polycystic ovarian syndrome prostaglandin progesterone receptor protein kinase A progesterone receptor knockout selective progesterone receptor modulator transforming growth factor β

INTRODUCTION Ovulation is a complex process with multiple physiological systems contributing to ensure the release of oocytes with the highest developmental potential at the

The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00014-5

appropriate time. Preovulatory ovarian follicles contain three main distinct cell lineages that arise through proliferation and differentiation as well as physical segregation (Fig. 1). The theca cells are specialized differentiated ovarian stromal cells separated from the granulosa layers by the follicular basement membrane, while two sublineages of granulosa cells are separated from each other by the follicular antrum. The granulosa cell subtypes diverge during follicular growth, through physical and biochemical influences arising from their close contact with the follicular wall (forming mural granulosa cells) or with the oocyte (cumulus cells). The physiological trigger for ovulation is the luteinizing hormone (LH) surge from the pituitary. In response to the LH surge, intrafollicular signals trigger the reactivation of oocyte meiosis, promote rapid production of a specialized extracellular matrix (ECM) leading to expansion of the cumulus oocyte complex (COC) and create a rupture pore in the follicular apex through which the COC is released. Thus, oocyte meiotic maturation is synchronized with release of the COC into oviduct for fertilization. In parallel, granulosa cells terminally differentiate into luteal cells which, in conjunction with neovasculogenesis and tissue restructuring, form the corpus luteum (CL). These processes are fundamental to successful establishment of pregnancy, but importantly also impact on the developmental potential of resultant embryos. Dramatic changes in gene expression mediate the reprogramming of granulosa, theca, and cumulus cell functions, with overlapping interdependent consequences in each of these compartments. Importantly, each process is also dependent on facilitative signaling from the oocyte to somatic cells, ensuring ovulation of healthy oocytes with full developmental competence for entry into the reproductive pool. Simultaneous with ovulation, the LH surge reinitiates meiosis so that the oocyte progresses to meiosis metaphase II (MII) and is ready for

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218 FIG. 1

14. OVULATION: THE COORDINATION OF INTRAFOLLICULAR NETWORKS TO ENSURE OOCYTE RELEASE

Intrafollicular events required for ovulation. The LH surge initiates multiple processes within the ovarian follicle including tissue remodeling, COC expansion, muscle contraction, and inflammation. The various layers and cell types within the follicle are also indicated.

LH surge Tissue remodeling

Cumulus cells Oocyte Muscle contraction

COC Expansion

Granulosa cells Basement membrane Inflammation

Theca interna/externa Tunica albuginea Surface epithelium

fertilization. In parallel, the uterus is prepared for implantation of the embryo through its response to rising progesterone released from the ovulating follicle. Thus, multiple aspects of female reproduction are initiated by the ovulatory LH surge and mediated through the response of the follicular somatic cells ensuring that fertilization potential and pregnancy readiness are tightly coordinated. Within the ovarian follicle, a number of intercellular events must be coordinated in order for ovulation to occur. Defects in any of these key events can result in anovulation and the entrapment of oocytes within a luteinized follicle. High circulating LH levels have a specific effect on the granulosa and theca cells and instigate a number of responses that are each essential for ovulation to occur (Fig. 1). (1) The cumulus cells rapidly synthesize a viscous ECM that enshrouds the oocyte and has a unique molecular composition essential for its release from the follicle. (2) The induction of proteolytic activity is necessary for follicular rupture and tissue morphogenesis including invasion of theca and vascular cells into the granulosa layer to form the CL. (3) Smooth muscle cells in the theca layer undergo contraction and cause thinning of the apical follicular wall. (4) Inflammatory mediators are generated, particularly prostaglandins and cytokines, which mediate an influx of immune cells into the ovary and surrounding the ovulating follicle. Thus, in general, the mural granulosa cells and theca cells respond first to the ovulatory trigger and induce a subsequent response in cumulus, vascular, and immune cells. Each of these cell types plays a critical role in eventual rupture and release of the oocyte and unique and tightly regulated patterns of gene and protein expression mediate the localization and temporal control of each of the key ovulatory events. This chapter will summarize our current understanding of the cellular events within the ovary that mediate ovulation. This commences with the intracellular

signaling that is activated in ovarian cells by surge levels of LH from the pituitary. Subsequently, each of the major intrafollicular processes that are essential for ovulation is described. These include cumulus-oocyte expansion, proteolytic and angiogenic tissue remodeling, muscle contraction, and inflammation and immune cell infiltration. The majority of studies revealing the specific gene products essential for ovulation have utilized mouse models; in particular, genetically modified mice lacking specific genes systemically (knockouts), or more recently mice lacking genes in specific cells within the ovary for instance in oocytes (GDF-9-Cre) or in granulosa cells (using Amhr2-Cre, Cyp19-Cre, or Pgr-Cre for temporal and spatially targeted gene deletions). Thus, the definitive studies that use mouse models to identify key ovulatory gene products are emphasized throughout. Lastly, clinical aspects of ovulation, namely anovulation and contraception are discussed.

OVULATORY INTRACELLULAR SIGNALING CASCADES The LH surge activates cognate LH receptors on theca and granulosa cells of mature follicles triggering activation of multiple intracellular signaling pathways that culminate in unique transcriptional complexes mediating expression of a suite of ovulatory genes (Fig. 2). Mural granulosa layers and theca cells of preovulatory follicles express LH-receptor (Lhcgr) mRNA and LH-R protein at levels typically an order of magnitude higher than cumulus cells in which Lhcgr expression is repressed by oocyte-derived signals [1,2]. Numerous studies indicate that preovulatory cumulus cells respond poorly, if at all, to direct LH exposure; reviewed in [3]. Interestingly, Lhcgr expression in thecal cells is regulated by the circadian clock gene Bmal1 [4] adding an element of diurnal

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temporal control to the ovulatory process. The theca specific LH-responsive, gene expression changes mainly relate to changes in steroid synthesis and response pathways [5]. The LH-R in granulosa cells (Fig. 2) is a classical Gαs-coupled G-protein-coupled receptor (GPCR) that activates adenylate cyclase, resulting in a large intracellular cAMP increase followed by cAMP-dependent serine protein kinase A (PKA) activation [6–8]. PKA phosphorylates the cAMP regulatory element-binding protein (CREB) which binds cAMP-response elements in specific gene promoters [9–11], then recruits the CBP/p300 transcriptional coactivators leading to activation of transcription [12,13]. Phosphodiesterases metabolize cAMP to also regulate intracellular cAMP tone and PDE4D, in particular, is an important regulator of cAMP levels in granulosa cells [14]. LH-R activation in granulosa cells also stimulates the extracellular regulated kinase (Erk1/2 or MAPK) pathway [15], with activation of Erk being very rapid [16] and dependent on PKA [11,15]. The activation of Erk in response to LH is necessary for ovulation [17]. The major targets downstream of Erk1/2 activation by the LH-surge are the CCAAT/Enhancer binding proteins alpha and beta (C/EBPα/β) which have been shown in knockout mouse studies to be highly important for ovulation [18]. The LH-R is also coupled to Gαq/11 protein complexes that lead to protein kinase C activation, which may specifically promote induction of progesterone receptor and subsequent key ovulation genes [19]. The Gαq/11/PKC

LH Gαs PDE4D

pathway likely activates AP1 transcription factors of the Jun, Fos, and Fra family which are also important in ovulation [20]. Thus several parallel signal transduction programs mediate the response to the surge in LH, with considerable overlap since PKA, PKC, and Erk kinases can all phosphorylate CREB and all three kinase pathways can also contribute to expression of critical genes, such as PGR [21] and C/EBPα/β in rodent models and humans.

Extracellular-Regulated Kinases Erk1/2 Extracellular-regulated kinases 1 and 2 [Erk1/2, also known as mitogen-activated protein kinases 3 and 1 (MAPK3/1)] are key regulators of cell proliferation and differentiation and participate in almost all LH-induced changes in the ovary at ovulation (Figs. 2 and 3). These signal transducing kinases are activated in granulosa cells through LHR-PKA directly as well as in cumulus cells via the action of granulosa cell-derived EGF-L interacting with EGF receptor on cumulus cells [22]. Mice null for Erk1 and with a granulosa-specific KO for Erk2 have profound and complete infertility, while mice with each mutation individually have normal fertility [17]. The follicles of the double mutant mice exhibit almost complete absence of the normal LH-induced responses; in fact, 77% of highly regulated LH target genes were dysregulated, including dramatically impaired expression of cumulus

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FIG. 2 Intracellular signaling activated by LH in granulosa cells. The LH surge activates adenylate cyclase (AC) and associated G-protein complexes to produce cAMP, activating protein kinase A (PKA) which phosphorylates the CREB transcription factor. CREB and transcriptional coactivators, that is, CBP/p300, recruited to DNA response elements induce ovulatory gene expression. PKA also activates extracellular regulated kinase (Erk1/2) which phosphorylates CEBPα/β transcription factors which and promotes complex formation with additional coactivators (CITED4, CBP) to induce ovulatory gene expression. One key CEBPβ dependent gene is progesterone receptor (PGR) which regulates essential ovulation genes. LH-R binding also activates protein kinase C (PKC) leading to AP1 family of transcription factor (Jun/Fos/Fra) activation and induction of ovulatory genes.

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FIG. 3

Gene products that are essential for ovulation. Luteinizing hormone (LH) binding to its receptor (LH-R) initiates a cascade of gene expression that controls the intrafollicular processes (expansion, proteolysis, contraction, and inflammation) required for ovulation. (1) Erk1/2 phosphorylates transcription factor Cited4 which together with CBP mediates the production of EGF-like ligands (EgfLs) by granulosa cells that are essential for induction of extracellular matrix (ECM) gene expression by cumulus cells. Cumulus matrix proteins integrate with serum proteins (such as IαI) to initiate cumulus expansion. Erk1/2 also phosphorylates transcription factor C/EBPb which (2) induces production of progesterone receptor (PGR) transcription factor which in turn induces a panel of genes essential for ovulation. These include protease Adamts1 which is required for follicular remodeling, and transcription factors (HIFs and PPARg) which induce endothelins (Edn2) which stimulate follicular smooth muscle contraction. (3) C/EBPb also induces expression of prostaglandin synthase 2 (Ptgs2) which generates prostaglandin E2 from free-fatty acid (ffa) precursors. PGE2 acts on receptors on cumulus cells (Pger2) to facilitate cumulus expansion and is also a key component of the inflammatory cascade that characterizes ovulation. (4) The LH surge initiates [via direct and indirect mechanisms (dashed line)] inflammatory responses throughout the follicle including nitric oxide synthase (NOS) expression in granulosa cells and leukocyte infiltration from the vasculature.

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and granulosa cell specific genes. As such, ovaries were profoundly affected with no cumulus expansion, no ovulation, absence of oocyte maturation, and no luteinization or progesterone production. These Erk1/2 mediated actions are mediated, at least partially, by induction of two transcription factors CEBPα and CEBPβ in granulosa cells [17,18] as well as CBP/p300-interacting transactivator with ED-rich tail (CITED-4) protein [10,13]. CITED4 forms a protein complex with CBP and C/EBPβ on the promoters of LH and Erk1/2 target genes, including Areg, Ereg, Btc, Ptgs2, Tnfaip6, Has2, Ptx3, Sulte1, and Pgr [13].

CCAAT Enhancer Binding Proteins CEBPα/CEBPβ CEBPβ, a transcription factor activated by Erk1/2 [17], is essential for ovulation since mice null for CEBPβ exhibit impaired ovulation [18,23]. Granulosa cell specific CEBPβ-null mice (CEBPβ gc-KO) have reduced expression of Ptgs2 and reduced luteinization genes (Star and

Contraction

Cyp11a1) post-hCG (Fig. 3). In cultured granulosa cells exogenous CEBPβ enables the induction of key genes Pgr, Tnaip6, Ptgs2, and Star [17] suggesting direct regulation. The anovulatory phenotype of the CEBPβ gc-KO is not as severe as the Erk1/2 gc-KO indicating that other Erk target genes are also involved in ovulation. One of these is likely to be transcription factor CEBPα. Similar to CEBPβ, CEBPα is induced within 2–4 h of the LH surge although CEBPα is downregulated in granulosa cells by 8 h while CEBPβ expression remains high in the CL [18]. CEBPαKO mice have a 30% reduction in ovulation in response to exogenous gonadotropins but interestingly have normal numbers of pups following natural mating possibly indicating a delay, rather than complete block in ovulation. Combining the CEBPαKO and the CEBPβ gcKO leads to a complete failure to ovulate similar to the Erk1/2 KO [18] suggesting that these two transcription factors together are key mediators of follicular rupture. Unlike the Erk1/2 KO however the CEBPα/ gcβKO have some degree of cumulus expansion, albeit it reduced, yet normal oocyte maturation with oocytes progressing to MII following hCG treatment. Microarray

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analyses of gene expression in granulosa cells demonstrate that Erk1/2 and CEBPα/β regulate mostly distinct gene cohorts. In general, Erk1/2 is essential for induction of early genes (such as Areg/Ereg, Pgr, Ptgs2, Ptx3) and CEBPs are essential for later genes in the ovulatory cascade, including genes required for luteinization, such as Cyp11a1, Lhcgr, and Star.

Progesterone Receptor Preovulatory granulosa cells begin producing progesterone in the days prior to the LH surge [7,24] whereas progesterone receptor (PGR) is only acutely induced after the LH surge [25] and immediately activated by the high local concentration of ligand. The nuclear progesterone receptor gene (Pgr) is among the most highly induced genes in ovarian granulosa cells reaching levels >100fold over basal expression within 4 h in granulosa cells responding to the LH surge, in rat and mouse [26], rhesus monkey [27,28], and humans [29]. A recent careful study of reproductive cyclicity in PGR-null rats confirmed its central importance in the ovary for ovulation and showed that PGR has a role, but is not essential for normal cyclicity [30]. Progesterone and its downstream targets are critical for follicular rupture and PGR is a highly specific regulator of this process. Mice with a targeted deletion of the Pgr gene (PRKO) exhibit normal follicle growth and luteinization, but a complete and specific block in ovulation [31,32]. Knockdown of PGR mRNA in primates [28] also results in a complete anovulatory phenotype. Similarly, antagonists of PGR, including RU486 [33] or more selective antagonists, such as ulipristal acetate [34–37], reduce or completely block ovulation; as do inhibitors of progesterone synthesis in species including rats [38], sheep [39], and humans [40]. Induction of Pgr expression requires adenylate cyclase/ cAMP/PKA-dependent signal transduction, and PKC activation adds a synergistic increase [41,42]. The β-catenin and Hippo family are also involved in Pgr regulation. Repression of Pgr by β-catenin and YAP1 in undifferentiated granulosa cells is released in response to the LH-surge [43,44]. The PGR nuclear receptor in turn regulates transcription of YAP1 target gene CTGF (connective tissue growth factor) leading to induction of known PGR targets, such as Adamts1 [45]. There are two main isoforms of PGR, of which the shorter form; PGR-A is predominant in rat granulosa cells [25] and is the most critical for ovulation, since mice lacking only PGR-A have severely reduced ovulation, while selective PGR-B ablation does not prevent ovulation [46]. PGR-A clearly induces genes that are critical effectors of ovulation; among which are two further transcription factors that mediate downstream ovulatory effects (Fig. 3). PPARg and HIFs (HIF1α, HIF1β, HIF2α)

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were identified as PGR-regulated genes via microarray [47–49]. PPARg is induced in mouse granulosa cells in response to hCG but not in PRKO mice [48]. When PPARg expression was inhibited by deleting the gene in periovulatory granulosa cells (using PGR-Cre), ovulation was reduced by almost 80%. Downstream target genes of PPARg were Et-2, cGKII, and IL6 (all previously identified as downstream of PR regulation) but not Adamts1. The ligand of PPARg in granulosa cells is not known but is possibly a COX-2 derived fatty acid since indomethacin inhibition of COX-2 blocked the induction of the panel of genes in a similar manner and this was reversed by the PPARg agonist rosiglitazone. [48]. Similarly, the HIF transcription factors are induced in granulosa cells in response to the LH surge but not in PRKO mice. Treating mice with a HIF inhibitor, echinomycin, concurrent with hCG dramatically inhibited ovulation [49] indicating that these transcription factors are essential mediators of ovulation. HIF target genes, again known to be downstream of PGR, include Edn2, Vegfa, and Cxcr4. These genes in particular may mediate vascular changes that contribute to ovulation, and of these Edn2 and Vegfa have been confirmed as pivotal for ovulation (see below). Thus, overlapping cascades of intracellular signals and gene induction within the periovulatory follicle (Figs. 2 and 3) are critical for normal ovulation and fertility. The cascade is set in motion by the surge of LH binding to LH-R which immediately induces PKA and PKC and Erk1/2 signaling. These subsequently activate a cohort of additional transcription factors including CEBPα/β, PGR, PPARg, and HIFs. These transcription factors in turn are responsible for induction of the functional effector processes that control ovulation. These can be grouped into four physiological responses: COC expansion, proteolytic activity and structural remodeling of the follicle, smooth muscle contraction in the thecal layer, and production of inflammatory mediators and immune cell infiltration.

CUMULUS OOCYTE COMPLEX EXPANSION Among the most striking and important events in ovarian follicles responding to the LH surge are the changes that occur in the COC, collectively referred to as “COC expansion”. This process includes the resumption of oocyte meiosis leading to extrusion of the first polar body as oocytes progress to MII (reviewed in accompanying chapter), an essential maturation step required for fertilization competence. In parallel, the COC mass expands many-fold in volume through a large induction of genes encoding extracellular proteins, proteoglycans and glycosaminoglycan synthesizing

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enzymes, resulting in formation of a specialized ECM within the cumulus layers (reviewed in Ref. [3]). In addition to expansion of their surrounding ECM, the cumulus cells themselves exhibit striking increases in adhesive capacity and migratory and invasive activity [50]. As cumulus cells are not directly responsive to the LH surge, COC expansion is activated via secondary paracrine signals (Fig. 3); namely the EGF-like ligands [amphiregulin (Areg), epiregulin (Ereg), betacellulin (Btc)] from granulosa cells [22] and prostaglandins from granulosa and cumulus cells [29,51,52]. Importantly, oocyte-derived factors are a required permissive signal for the specific cumulus gene expression [53,54]. This somatic cell-oocyte dialogue synchronizes the activation of oocytes that have acquired developmental competence with release from the follicle, and the process is coordinated with the endocrine LH cycle by the granulosa cells.

Extracellular Matrix Production The rapid expansion of the COC in periovulatory follicles involves the production of a specific suite of ECM proteins most of which have been shown to be important for ovulation in animal models, indicating that COC expansion is a critical ovulatory event. The backbone of the expanded cumulus matrix is hyaluronan (HA), a large disaccharide chain common to many extracellular matrices including cartilage where it generates viscoelastic properties, and evidence suggests it has similar influence in the COC [55]. Synthesis of HA utilizes glucose as a substrate for synthesis of glucosamine and glucuronic acid via the hexosamine biosynthesis pathway. The hyaluronan synthase enzyme HAS2, induced in cumulus cells by EGF-L, FSH, and prostaglandins assembles the HA subunits into very long disaccharide HA chains (up to megadaltons) and extrudes it into the extracellular space [56]. Many HA-interacting proteins are also induced in cumulus cells through the same signaling pathways. Interalpha trypsin inhibitor (IαI) is a circulating protein complex secreted by the liver and which enters the follicle due to heightened vascular permeability in periovulatory follicles [57,58]. Heavy chains (HCs) of IαI form a covalent bond with HA and organize the specific structure and functions of the COC matrix. Disruption of the ability of IαI HC to bind HA results in 2% of all genes expressed in arterial endothelial cells are regulated by HIF1, directly or indirectly [39].

Luteal HIF1A-Dependent Genes Different steps of angiogenesis are promoted following hypoxic challenge [40, 41] (see Fig. 1). Reports on inactivating mutations of the HIF pathway have emphasized the importance of HIF1A in regulating genes involved in angiogenesis [15, 16, 39], in fact, substantial number of these genes contain HREs in their promoters [41, 42]. Vascular endothelial growth factor A (VEGFA) occupies a central place among the aforementioned genes. Also known as vascular permeability factor, VEGFA is the major specific stimulator of endothelial cell proliferation and migration, acting through two tyrosine kinase receptors, VEGFR-1 (flt-1) and VEGFR-2 (KDR; [43]). In hypoxia, VEGFA transcription is upregulated by HIF [40]. Hypoxia also leads to stabilization of VEGFA mRNA [44]. The intrinsically short half-life of VEGFA mRNA (approximately 30 min) is significantly extended under stress, presumably through hypoxia-augmented binding of one or more unidentified proteins to its 30 untranslated region (30 UTR) [45]. The biological activity of secreted

VEGFA is further influenced by the hypoxia-inducible expression of the Flt-1 and KDR receptors [46, 47]. In the CL, VEGFA expression is upregulated soon after the LH surge and elevated VEGFA levels (mRNA and protein) remain at least until the mid-to-late luteal phase [13, 23, 24, 48]. In most species, VEGFA mRNA is detected in new CL in the granulosa-derived lutein cells [48, 49]. However, a study by Redmer et al. (2001) demonstrated that it is also expressed in pericytes in ovine CL [50]. Several studies had verified that VEGFA is essential for optimal CL function [21, 51, 52]. Fraser et al. studied the role of VEGF Trap (a recombinant, chimeric protein comprising Ig domains of human VEGF-R1 and R2 [52]) on luteal function. A single injection of VEGF Trap administered to macaques shortly after ovulation, blocked the normal luteal phase elevation of progesterone. In addition, in marmoset monkeys, pharmacological inhibition of VEGFA (using the VEGFA antibody aflibercept) suppresses luteal progesterone and prevents the successful establishment of pregnancy [53], demonstrating that VEGFA is essential for both the development and maintenance of normal luteal function in primates. Similarly, daily intraluteal injections with VEGFA antibody (from day 1 to day 8) markedly decreased the bovine CL volume and plasma progesterone concentrations [51]. As in other tissues, hypoxia (via HIF1A) is also a strong inducer of VEGFA in the CL. In luteinized granulosa cells cultured under hypoxic conditions (low O2) or hypoxia-mimicking agents such as Cobalt chloride (CoCl2), it stimulates VEGFA [25, 54–56]. HIF1 activity in luteal cells transfected with oligodeoxynucleotides containing an HIF1-binding site, reduced VEGFA mRNA levels under hypoxia, as did HIF1A silencing [25, 54–56]. These results further support the notion that HIF1 mediates the transcriptional activation of the VEGFA gene in the CL (Fig. 1).

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Interestingly, despite the evidence previously mentioned, no overlap exists between VEGFA and HIF1A profiles during CL development [23, 57]. The presence of HIF1A protein is limited to the very early stages of CL development, whereas high VEGFA progresses to later stages of the CL lifespan, suggesting the involvement of additional factors (Fig. 2). Endothelin-2 (EDN2) is a new player in reproductive processes implicated in ovulation and CL formation [54, 58, 59]. The multiple effects of EDN2 are mediated by two G-protein-coupled receptors, termed ETA and ETB, exhibiting distinct molecular and pharmacological characteristics [60]. Prevention of EDN2 action in rodents using EDN receptor antagonists resulted in unruptured follicles that failed to develop into CL [58, 59, 61]. Consistent with the contention that EDN2 is essential for CL formation, it was reported that EDN2 mRNA levels were transiently expressed immediately after ovulation, in early, developing bovine CL [54] see Fig. 2. EDN2 in follicles or CL was localized mainly to the luteinized granulosa cells or luteal steroidogenic cells, respectively [54, 61, 62]. Supporting the role of EDN2 in ovulation it was reported that granulosa cells collected from patients with polycystic ovary syndrome (PCOS), characterized among other symptoms, by chronic oligo-ovulation or anovulation, had significantly lower EDN2 mRNA expression when compared with normally ovulating women [62]. EDN2 was identified as a hypoxia-regulated gene [54, 61]. In fact, hypoxia was found to be a strong inducer of EDN2 transcription in granulosa-lutein cells of all species examined thus far (rat, mice, and cows; [25, 54, 63]). Low oxygen or the presence of CoCl2 led to the simultaneous elevation of HIF1A and HIF2A proteins and EDN2 in granulosa cells [25, 54, 56, 64]. Moreover, HIF1A knockdown with specific siRNA or the addition of echinomycin, which interferes with the binding of HIF1A/B heterodimers, abolished hypoxia-induced EDN2, thus confirming that EDN2 is a HIF1A-responsive gene [25, 56, 65]. A putative HRE sequence between 613 and 629 with a core similarity of 0.9 was identified while exploring the human EDN2 promoter (1500 bp) using MatInspector Genomatix software [66]. This site could mediate the induction of EDN2 by hypoxia, although promoter studies are still needed to validate this contention. HIF1 is also known to induce the expression of glucose transporter 1 (GLUT1) [67]. GLUT1 mRNA expression in bovine CL was higher at the early luteal stage compared with the other later stages, and hypoxia increased GLUT1 mRNA expression in luteal cells [68]. GLUT1 inhibition decreased progesterone production, whereas glucose supplementation elevated progesterone. These results therefore suggest that GLUT1 (induced by hypoxic conditions in the early CL) plays a role in establishing and developing bovine CL, by supporting luteal progesterone synthesis at the early luteal stage.

There are other HIF1-regulated genes that are tentatively involved in angiogenesis, such as nitric oxide synthase (NOS3; [69] and heme oxygenase 1 [70]. Hypoxia-Independent Regulation of HIF1A HIF1A can also be hormonally regulated in an oxygenindependent manner. Insulin-like growth factor 1 (IGF1), for instance, augmented the accumulation of HIF1A in normoxia in several cell types [71]. Reactive oxygen species (ROS) is another factor implicated in HIF1A accumulation [56]. Evidence suggests that ROS, paradoxically, increases HIF1A by directly inhibiting PHD catalytic activity [72, 73]. Yalu et al. examined the involvement of ROS using hydrogen peroxide (H2O2) and a broadrange ROS scavenger (BHA). They reported that H2O2 elevated the levels of HIF1A protein together with elevated EDN2 and VEGFA mRNA levels in human granulosa lutein cells (Fig. 1). Furthermore, BHA significantly reduced the levels of HIF1A protein induced by CoCl2 and the expression levels of HIF1A-dependent genes (EDN2 and VEGFA) were markedly reduced [56]. However, due to its role in CL formation and luteinization, the involvement of LH/hCG canonical signaling pathway-cAMP—in the normoxic induction of HIF1A remains a highly physiologically significant issue. There is a strong evidence that both VEGFA and EDN2 within the CL are regulated not only by hypoxia but also by LH/hCG. VEGFA expression in human and bovine luteinized granulosa cells was shown to be dose- and time dependently enhanced by hCG in vitro [23, 54, 74, 75]. Furthermore, bolus injection of hCG to macaques elevated VEGFA levels in follicular fluid [76]. Similarly, LH induced EDN2 mRNA in luteinized human and bovine granulosa cells [54, 56]. Do gonadotropins directly augment the transcriptional activity of HIF1A under normoxic conditions? And do LH/hCG and hypoxia signaling synergize in elevating HIF1A activation? Both LH and FSH [77, 78] as well as forskolin [25, 64] elevated VEGFA and EDN2 without any apparent increase in HIF1A protein. However, Rico et al. [78] provided evidence demonstrating that gonadotropinregulated Vegfa requires HIF1A transcriptional activity. They reported that granulosa cells from mice that lack a single HRE in the Vegfa promoter failed to respond to FSH or LH with an increase in Vegfa mRNA [78]. Shrestha et al. [64] provided additional evidence that elevated cAMP utilizes the HIF1A pathway for inducing the hypoxia-dependent gene, EDN2. They observed that under normoxic conditions, forskolin (adenylyl cyclase activator) triggered changes typical of hypoxia. It elevated HIF1A and EDN2, whereas HIF1A silencing greatly reduced forskolin’s ability to elevate EDN2. Treatment with hCG or cAMP analogues promoted HIF1A mRNA [21, 56, 79], but this did not translate into

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INTRODUCTION

appreciably higher HIF1A protein levels [23, 79]. Similarly, using bovine granulosa cells, it was found that LH elevated the mRNA levels of HIF1A without affecting its protein concentration [56]. However, there was a synergistic, dose-dependent effect of LH (or forskolin) with CoCl2 on HIF1A protein levels [56]. The combined effect results from HIF1A being induced transcriptionally by LH and posttranscriptionally by hypoxia-induced stabilization. Importantly, the synergism between cAMP on HIF1A was also manifested in higher VEGFA and EDN2 mRNA levels [64, 78] (Fig. 1). Role of microRNA-210 in Hypoxic Responses Several hypoxia-responsive microRNAs (miRs) have been identified [80].MiRs are short, small noncoding RNAs; they regulate gene expression at the posttranscriptional level by either degradation or translational repression of target mRNAs regulating physiological and pathological processes [81]. Among these miRs, the most consistent and powerful response to hypoxia is observed in miR-210. Because of its robust response to hypoxia and its role in mediating multiple physiological processes, miR-210 has been termed the “master hypoxamir.” The increased expression of miR-210 is linked to an in vivo hypoxic signature. MiR-210 is a direct transcriptional target of HIF1A [82–84]. MiR-210 can simultaneously regulate the expression of multiple target genes in order to finetune the adaptive response of cells to hypoxia. MiR-210 also participates in the response to oxygen deprivation and plays a role in the regulation of angiogenesis. For instance, it was shown that miR210 regulates the expression of both VEGFA and VEGFR2 and contributes to hypoxia or ischemic diseases [85]. The report of Shrestha et al. highlighted the role of miR-210 as an important component of the adaptive response to HIF1A in human granulosa-lutein cells [64]. miR-210 was elevated in those cells exposed to hypoxia, whereas HIF1A silencing prevented this induction [64]. The levels of miR-210 were positively correlated with the hypoxia-responsive genes in these cells, namely, EDN2. Elevating endogenous miR-210 by hypoxia and miR-210-mimic increased EDN2, whereas inhibiting miR-210 reduced EDN2 even in the presence of CoCl2, implying the significance of miR-210 in the hypoxic induction of this gene [64]. In support, it was recently shown that miR-210 was differentially expressed during follicular-luteal transition in buffalo ovary [86]. In addition, it was recently reported that miR-210 was drastically upregulated in early CL compared with other stages of the CL cycle [87]. As for miR’s other function, miR-210 downregulates its target mRNAs in a sequence-specific way. Many miR-210 gene targets were identified, two of which, glycerol-3-phosphate dehydrogenase 1-like (GPD1L) and succinate dehydrogenase complex subunit D (SDHD),destabilize HIF1A [88, 89].

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Transfecting human granulosa lutein cells with miR210-mimic (elevating miR-210 levels by >500-fold) markedly suppressed GPD1L and SDHD, implying that these genes are miR-210 targets [64]. Pertinent to miR-210’s role in hypoxia, these two genes were shown to destabilize HIF1A, but with different modes of action [88, 90]. Reduced SDHD induced succinate accumulation, a Kreb cycle intermediate that is a natural inhibitor of PHD [89, 90], thus stabilizing HIF1A. GPD1L regulates HIF1A differently; high amounts of GPD1L caused increased proline hydroxylation of HIF1A [88], resulting in proteosomal degradation of HIF1A protein [26]. Less vigorous miR-210 induction in hypoxia could still reduce GPD1L but much less so SDHD; suggesting that higher concentrations of miR-210 may be necessary to suppress SDHD in granulosa cells [64]. Experiments utilizing either hypoxia-induced miR-210 or miR-210 overexpression suggest that GPD1L reduction plays a role in EDN2 expression. GPD1L silencing by its siRNA provides direct evidence for this assumption, showing increased HIF1A protein and elevated EDN2. Taken together, these data imply the need to suppress GPD1L, by elevated miR210, in order to allow HIF1A accumulation and an increase in EDN2. Forskolin, although less prominently, also elevated miR-210 and EDN2 while reducing GPD1L, suggesting that forskolin initiates a response similar to that of hypoxia. Furthermore, similar to hypoxia, most of these effects of forskolin were abolished with HIF1A silencing, demonstrating the significance of HIF1A/ miR-210 also in cAMP-elevated EDN2 (Fig. 1). These in vitro findings suggest that the two main signals driving CL formation, namely, hypoxia and LH (mimicked in vitro by forskolin) act similarly via the HIF1A/miR-210/GPD1L loop to augment EDN2 expression, as illustrated in Fig. 1.

Luteal Angiogenesis The formation of a well-developed capillary bed in this gland is indispensable for establishing a suitable microenvironment for differentiation and maturation of a fully functional CL [2, 4, 15, 91]. The short period of angiogenesis (until day 5 of the cycle in the cow) is later followed either by maintenance and stabilization of the vasculature, such as that occurring during pregnancy, or the controlled regression of the microvascular tree in the nonfertile cycle during luteolysis [5]. It has been estimated that in the mature CL up to 50% of cells are of vascular origin [2, 4, 15, 91]. Owing to this extensive vascularization, the CL receives one of the highest blood supplies, per gram of tissue, of any organ [92]. The intense vascularization of the CL during the midphase of the estrus cycle allows an extremely high rate of blood flow, which enables elevated progesterone perfusion;

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therefore, progesterone concentrations in the systemic circulation are highly correlated with luteal vasculature density. Early capillary sprouts, which originate from venules of the former theca interna, invade the folds of granulosa cells [93]. In bovine and sheep, capillary sprouts are evident shortly after ovulation, 36 h in bovine [93] and in rats, as early as 12 h postovulation [94]. Endothelial cell migration takes place only after the basal lamina, which separates the avascular stratum granulosum from the theca interna, has been enzymatically broken down. Immediately after ovulation, the tissue folds of the collapsed follicle are composed of granulosa cells with morphological signs of initial luteinization. Not only endothelial cells, but also other cell types, such as eosinophils (eosinophilic granulocytes and globular leukocytes) and macrophages, migrate from the surrounding ovarian stroma in great numbers in the developing CL [95], as depicted in Fig. 3. An increased number of eosinophils invade the preovulatory follicles shortly before ovulation in ovaries of several species [95]. Eosinophils may contribute to the angiogenic process by releasing numerous growth factors, cytokines, and chemokines, most prominently, VEGFA and interleukin 18, which enhance proliferation and serve as chemoattractive agents for endothelial cells [95, 96]. Treatment with dexamethasone not only suppressed eosinophilic cells migration into the developing CL, but also caused a significant reduction in progesterone levels in the mid and

FIG. 3 Electron micrograph of bovine ovary 1 day postovulation. Eosinophilic granulocyte immigrated from the blood vessels of the theca interna one day after ovulation. EC, immigrating endothelial cells; EOG, eosinophilic granulocyte; ER, erythrocyte.

late luteal phases (days 8–17) [95]. A similar decline in progesterone was also found in the CL of prednisolonetreated sheep [97]. As the CL matures, the proliferation rate of endothelial cells is reduced, compared with the developing CL. The well-developed vasculature becomes stabilized in mature CL. Arteries running in the peripheral zone of the CL, which arise from a single, spiral artery, break up into septal arteries. They run in the septal connective tissue, which separates luteal tissues, and penetrate the parenchyma. Finally, the parenchymatous arteries split up and form a dense capillary network between the luteal cells. As in many other endocrine glands, these capillaries are lined by a fenestrated endothelium. It has been shown that most luteal cells are adjacent to one or more capillaries, which certainly enhances the intense interactions between luteal cells and endothelial cells; however, ultrastructural observations have indicated that no immediate contact exists between the luteal cells and the endothelial cells [93]. Proangiogenic Factors: VEGFA, FGF2, and PGE2 VEGFA, as discussed before, is the major specific stimulator of endothelial cells’ proliferation and migration in the CL. Along with VEGFA, fibroblast growth factor 2 (FGF2) provides essential proangiogenic support for the developing CL [57, 98, 99]. FGF2 is abundant in the CL; in fact, FGF2 was first extracted and identified in bovine CL [100]. FGF2 stimulates the proliferation of endothelial cells derived from bovine CL [101, 102].FGF2 also acted as survival factor for luteal endothelial cells, by suppressing caspase-3 activation alone and inhibiting the expression and function of thrombospondin1, a known apoptotic factor [103]. Its importance became evident by experiments showing that local neutralization of FGF2, by directly injecting FGF2 antibody into the developing bovine CL, altered luteal growth and function, probably by inhibiting the establishment of a new vascular network during CL formation [51]. In general, FGF2 is more potent at inducing endothelial cell proliferation than is VEGFA. Indeed, treatment with an FGF receptor1-signaling inhibitor (PD173074) almost completely blocked the formation of luteal endothelial networks in vitro [99]. The effects of the FGF receptorsignaling inhibitor occurred even in the presence of exogenous VEGFA, which suggests that FGF2 is critical for forming luteal endothelial networks [99, 101]. FGF2 levels, like those of VEGFA, are elevated in the young CL; however, while VEGFA is maintained for the most of CL’s lifespan, FGF2 peaks at a very early luteal phase [57]. This early rise of FGF2 is most probably a result of the LH, as demonstrated by in vitro studies using luteal granulosa cells [57, 103] and by ovulation induction experiments in cows demonstrating that FGF2 is temporally induced by LH surge [104] see in Fig. 2.It is well

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known that the ovulatory surge of LH also induces the rate-limiting enzyme in prostaglandin synthesis (PTGS2) and prostaglandin endoperoxide 2 (PGE2) in granulosa cells [105] see in Fig. 2. This increase is an essential component of the ovulatory process as demonstrated in numerous studies (see for example [106–108]). Role of PGE2 in Luteal Angiogenesis There is increasing evidence that PGE2 is an important stimulator of angiogenesis accompanying CL formation [109–111]. Suppressed vasculature formation was observed in newly formed rat CL, and in primate ovulatory follicles treated with PTGS2 inhibitors, replenishing PGE2 alleviated by this inhibition [110, 111].PTGS2 is abruptly induced in response to LH in the granulosa cells of follicles approaching ovulation [105, 112, 113].PGE2 action is mediated by its four distinct receptors: PTGER1, PTGER2, PTGER3, and PTGER4 [114]. PTGER1 and PTGER2 agonists promoted monkey and human ovarian microvascular endothelial cells migration and sprout formation [109, 110]. Likewise, PTGER2 antagonist in rats suppressed luteal endothelial cells tube formation [115] emphasizing the importance of PGE2 in luteal angiogenesis. A potential connection between PGE2 and FGF2 had also been reported, FGF2 knockout or FGFR1 blockade impaired the ability of PGE2 to induce in vitro angiogenesis in aortic endothelial cells [116, 117]. The interesting possibility of PGE2 acting directly to stimulate FGF2 in granulosa cells, as additional mechanism to support angiogenesis in the CL, remains to be explored.

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The ECM surrounding the capillary sprouts contains a delicate network of fine fibrils consisting of fibronectin, which forms a wide-mesh network around the capillary sprouts [93]. Another ECM protein, type IV collagen, also forms a fine and discontinuous layer around the capillary sprouts. In the ECM of young capillary, sprouts are patchy and amorphous. In more developed sprouts however, staining for collagen type IV and laminin displayed a distinct immunopositive line around endothelial cells and pericytes. These observations indicate that the ECM surrounding the newly forming microvessels undergoes a series of changes before achieving a final maturation. Ultrastructural investigations in the bovine luteinizing tissue on days 1 and 2 postovulation revealed that early capillary sprouts were usually preceded by pericytes migrating at the tip of the sprouts [93, 122]. In fact, no pericyte-free endothelial sprouts could be found, supporting the idea that pericytes play an essential role as leading structures for capillary sprouting [93] (see Fig. 4).

Matrix Remodeling The process of angiogenesis involves a series of events that largely depend on proteinases and their ability to remodel the ECM, starting with degradation of the basement membrane to allow for endothelial cell breakthrough, migration, and proliferation. This is followed by organization into nascent blood vessel sprouts, vessel maturation and stabilization, deposition of basement membrane around the new vessels, and finally pruning or remodeling of the new vasculature for physiological needs [118]. Likewise, the formation of the CL and the angiogenic processes are accompanied by dramatic tissue remodeling that requires controlled and targeted proteolysis of ECM and basal lamina components [119–122]. The stromal cells of the theca interna and theca externa are incorporated into the centers of the folds and the margins of the developing CL. Since the ECM also acts as a reservoir for paracrine and endocrine signals, where FGFs are notable examples, ECM remodeling within the CL adjusts its bioavailability within the gland [123].

FIG. 4

Schematic representation of early angiogenic processes in the forming CL. Endothelial cells (EC) form sprouts from existing capillaries and venules under the guidance of and in close association with pericytes (PC). These early angiogenic events can be subdivided into several steps: 1. Partial dissolution of the basal membrane (BM) of the vascular wall. 2. Migration and elongation of PC and EC at the leading tip of the sprout. 3. Immediate canalization of the endothelial sprout. 4. Mitosis (M) of ECs behind the leading front of migrating ECs and PCs. 5. Further differentiation of the new capillary and rebuilding of a fully developed BM.

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Amselgruber and his colleagues [93] also showed slender cellular structures between pericytes and endothelial cells, which penetrate deeply into the neighboring cells. These two cell types are embedded within the same basal lamina produced by the two cell types [124]. In addition to their guiding role in the sprouting process [4, 93], pericytes are important producers of VEGFA, the production of which is stimulated by endothelialderived nitric oxide [4]. It has been convincingly demonstrated that a link exists between the expression of angiogenic factors and the activity of matrix-degrading proteases, especially Matrix Metalloproteinases (MMPs) [122]. MMPs constitute a family of structurally related proteins that degrade ECM and basal laminae. They degrade the basement membrane and the nearby ECM surrounding established blood vessels at the onset of angiogenesis, and release proangiogenic growth factors that otherwise would remain bound to the ECM. The activity of MMPs is balanced by their inhibitors, the tissue inhibitor of matrix metalloproteinases (TIMPs) [119, 120]. The plasminogen activator (PA) is another important proteolytic system involved in the control of fibrin degradation, matrix turnover, and cell invasion. The components of the PA system include urokinase-plasminogen activator (uPA) and tissue-type plasminogen activator (tPA), together with the major inhibitors of this system: plasminogen activator inhibitor-1 and -2 (PAI-1, PAI-2, SERPINES) [125]. The mRNA expression of MMP9 significantly increased, ranging from its levels in follicle to CL tissue, with the highest expression on days 8–12. MMP19 showed a comparable mRNA expression profile, with an immediate increase after ovulation [122]. Whereas tPA displayed constant expression during the late follicular and the early to midluteal phase, uPA, its receptor (uPAR), as well as PAI-1 and PAI-2 levels all rose from the follicular phase onwards up to the midluteal phase [122]. Studies in the rat [126] and rhesus monkey [127] found a distinct localization of uPA and PAI-1in the developing CL. uPA was localized to the endothelial cells along the route of capillary extensions, whereas PAI-1 was expressed in fibroblasts near the capillary sprouts. PAI-1, TIMP-1, and TIMP-2 are also produced by large ovine luteal cells [121]. PAI-1 may protect neovascularized tissue from excessive proteolysis [126]. Interactions between uPA and its receptor appear to be mandatory for the angiogenic effect of uPA [128]. Owing to the activity of matrixdegrading enzymes, endothelial cells in the CL-expressing uPA/uPAR interact with PAI-1 secreted from fibroblasts, become detached from the ECM, and can migrate into the unvasculated stroma to form new capillaries [95]. The composition of the ECM between the luteal cells and the vasculature has been reviewed in great detail by Irving-Rodgers et al. (2006, 129). Collagens I, III, and IV, laminins, various chondroitin sulfate proteoglycans,

such as versican, and high-molecular-weight linear polysaccharides, such as hyaluronan, have been localized by different species to the ECM and may be essential for maintaining the luteal cell phenotype [129].

CONCLUSIONS CL formation entails, at a cellular level, intense interactions between microvascular cells (endothelial cells and pericytes), steroidogenic cells, and migrating immune cells, leading to cell differentiation, angiogenesis, and ECM remodeling. The process of CL development bears many similarities to other physiological or pathological tissue formation. It involves rapid growth with initially unmatched vascularization leading to hypoxic conditions, which eventually promote angiogenesis and cell metabolism. The angiogenic process accompanying the formation of the CL or tumor requires, in both cases, tissue remodeling that involves controlled proteolysis of ECM and basal lamina components [118–122]. In accordance, VEGFA plays a key role in both CL [3] and tumor development [40, 46]. EDN2 affects early CL development [54, 56] but it was also identified as a promoter of cancer progression [130–132]. Furthermore, elevated HIF1A and miR-210 levels characterize early CL as well as cancer development [27, 133]. However, whereas CL maintenance is tightly controlled, cancer cells continue to proliferate and tissue growth evades regulation. The principal difference between CL and tumor in this respect is the hormonal control over CL formation and maintenance. The functional cells of the CL, the steroidogenic cells, gradually differentiate in response to LH and their proliferation is halted. Interestingly, while HIF1A, VEGFA, EDN2, and miR-210 are regulated almost exclusively by lack of oxygen in cancer cells, in the CL they have adopted additional distinct regulatory mechanism, i.e., LH and its main signaling molecule, cAMP [56, 64]. Moreover, the CL lifespan is not only controlled by LH acting as a luteotrophic hormone, but also by PGF2A terminating the function and mere presence of CL [5]. Whether these lessons can be applied to control tumor progression remains to be elucidated.

Glossary Angiogenesis The growth of new blood vessels from the existing vasculature. cAMP A second messenger, derivative of adenosine triphosphate (ATP). Chondroitin sulfate proteoglycans Proteoglycans consisting of a protein core and a chondroitin sulfate side chain. Endothelin-2 (EDN2) A small peptide, expressed and is mainly localized to the granulosa cells within the follicle, as observed in bovine, murine, and human ovarian tissue. Endothelin-2 has functions within the ovarian follicle related to ovulation. Fibroblast growth factor 2 (FGF2) A growth factor and signaling protein that influences broad mitogenic and cell survival activities.

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Proangiogenic factor. Also known as basic fibroblast growth factor (bFGF) and FGF-beta. Folliculo-luteal transition The transition phase in between follicular phase and luteal phase of the reproductive cycle initiated by the surge in LH accompanying ovulation and formation of corpus luteum. Glycerol-3-phosphate dehydrogenase 1-like (GPD1L) Catalyzes the reversible biological conversion of dihydroxyacetone (DHAP) to glycerol-3-phosphate. A novel regulator of HIF-1alpha stability and a direct target of miR-210. Hypoxamir (miR-210) A short RNA molecule that is upregulated in response to hypoxia-inducible factors. Hypoxia Low oxygen tension. Hypoxia response elements (HRE) The binding site for hypoxiainducible factors in the promoter regions of hypoxia-inducible target genes. Hypoxia-inducible factor 1 (HIF1) The transcription factor, composed of a heterodimer of HIF-1alpha and HIF-1beta subunits, which functions as a global regulator of O2 homeostasis and the adaptation to O2 deprivation. Luteal insufficiency A condition that occurs when the luteal phase is shorter than normal, progesterone levels during the luteal phase are below normal, or both which interferes with the implantation of embryos. Polycystic ovary syndrome (PCOS) The most common endocrine disorder among women due to set of symptoms such as oligo-ovulation, high androgen levels, insulin intolerance, and ovarian cysts. Prostaglandin Endoperoxide 2 (PGE2) A key prostaglandin-mediating ovulatory process. Induced by LH in follicular granulosa cells. Prostaglandin F2a (PGF2a) The natural luteolysin. Released from the endometrium and terminates the CL life span and progesterone production. Reactive oxygen species (ROS) A chemically reactive chemical species containing oxygen, which is formed as a natural byproduct of the normal metabolism of oxygen and has important roles in cell signaling and homeostasis. Increase in ROS levels can cause oxidative stress. Tissue-type plasminogen activator (TPA) A serine protease that converts inactive plasminogen to plasmin and promotes degradation of fibrin. Hormonally induced mainly in granulosa cells and oocyte. Type IV collagen The main collagen component of the basement membrane forming a network that underlies epithelial and endothelial cells and functions as a barrier between tissue compartments. Vascular endothelial growth factor A (VEGFA) A mitogen that specifically acts on endothelial cells and has various effects, including mediating increased vascular permeability, inducing angiogenesis, vasculogenesis and endothelial cell growth, promoting cell migration, and inhibiting apoptosis.

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[97] Murdoch WJ, Van Kirk EA. Aetiology of attenuated luteal development in prednisolone-induced eosinopenic ewes. Reprod Fertil Dev 2000;12(3–4):127–32, 11302421. [98] Gabler C, Plath-Gabler A, Killian GJ, Berisha B, Schams D. Expression pattern of fibroblast growth factor (FGF) and vascular endothelial growth factor (VEGF) system members in bovine corpus luteum endothelial cells during treatment with FGF-2. VEGF 2004;39(5):321–7, 15367264. [99] Woad KJ, Hammond AJ, Hunter M, Mann GE, Hunter MG, Robinson RS. FGF2 is crucial for the development of bovine luteal endothelial networks in vitro. Reproduction 2009;138(3):581–8, 19542253. [100] Gospodarowicz D, Cheng J, Lui GM, Fujii DK, Baird A, Bohlen P. Fibroblast growth factor in the human placenta. Biochem Biophys Res Commun 1985;128(2):554–62, 3994712. [101] Woad KJ, Hunter MG, Mann GE, Laird M, Hammond AJ, Robinson RS. Fibroblast growth factor 2 is a key determinant of vascular sprouting during bovine luteal angiogenesis. Reproduction 2012;143(1):35–43, 21998077. [102] Zalman Y, Klipper E, Farberov S, Mondal M, Wee G, Folger JK, et al. Regulation of angiogenesis-related prostaglandin f2alphainduced genes in the bovine corpus luteum. Biol Reprod 2012;86(3):92, 22174022. [103] Farberov S, Meidan R. Functions and transcriptional regulation of thrombospondins and their interrelationship with fibroblast growth factor-2 in bovine luteal cells. Biol Reprod 2014;91(3):58, 25061096. [104] Berisha B, Steffl M, Amselgruber W, Schams D. Changes in fibroblast growth factor 2 and its receptors in bovine follicles before and after GnRH application and after ovulation. Reproduction 2006;131(2):319–29, 16452725. [105] Sirois J, Sayasith K, Brown KA, Stock AE, Bouchard N, Dore M. Cyclooxygenase-2 and its role in ovulation: a 2004 account. Hum Reprod Update 2004;10:373–85, 15205395. [106] Peters MW, Pursley JR, Smith GW. Inhibition of intrafollicular PGE2 synthesis and ovulation following ultrasound-mediated intrafollicular injection of the selective cyclooxygenase-2 inhibitor NS-398 in cattle. J Anim Sci 2004;82:1656–62, 15216991. [107] Davis BJ, Lennard DE, Lee CA, Tiano HF, Morham SG, Wetsel WC, et al. Anovulation in cyclooxygenase-2-deficient mice is restored by prostaglandin E2 and interleukin-1beta. Endocrinology 1999;140(6):2685–95, 10342859. [108] Richards JS. Ovulation: new factors that prepare the oocyte for fertilization. Mol Cell Endocrinol 2005;234(1–2):75–9, 15836955. [109] Trau HA, Brannstrom M, Curry Jr. TE, Duffy DM. Prostaglandin E2 and vascular endothelial growth factor A mediate angiogenesis of human ovarian follicular endothelial cells. Hum Reprod 2016;31(2):436–44, 26740577 Pubmed Central PMCID: 4716810. [110] Trau HA, Davis JS, Duffy DM. Angiogenesis in the primate ovulatory follicle is stimulated by luteinizing hormone via prostaglandin E2. Biol Reprod 2015;92(1):15, 25376231 Pubmed Central PMCID: 4434930. [111] Sakurai T, Tamura K, Kogo H. Stimulatory effects of eicosanoids on ovarian angiogenesis in early luteal phase in cyclooxygenase-2 inhibitor-treated rats. Eur J Pharmacol 2005;516(2):158–64, 15921676. [112] Berisha B, Schams D, Rodler D, Sinowatz F, Pfaffl MW. Changes in the expression of prostaglandin family members in bovine corpus luteum during the oestrous cycle and pregnancy. Mol Reprod Dev 2018;29877057. [113] Tsai SJ, Wiltbank MC, Bodensteiner KJ. Distinct mechanisms regulate induction of messenger ribonucleic acid for prostaglandin (PG) G/H synthase-2, PGE (EP3) receptor, and PGF2 alpha

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C H A P T E R

17 Luteolysis and the Corpus Luteum of Pregnancy Camilla K. Hughes, Joy L. Pate Abbreviations ABCA1 ACAT1 AKR1C1 AKR1C3 AKT AMPK ANGPT ATF3 ATG CAMKK2 cAMP CASP3 CBR1 CCL CD CSH CXCL CYP11A1 DAG DDX58 EDN1 EGR1 ESR1 FAS FGF2 FOS FOXP3 GATA3 hCG HSD3B1 IDO IFIH1 IFN IL IP3 ISG ISG15 JAK-STAT JUN LDLR

ATP-binding cassette subfamily A member 1 acetyl CoA acetyl transferase 1 aldo-keto reductase family 1 member C1 (also known as 20-alpha-hydroxysteroid-dehydrogenase) aldo-keto reductase family 1 member C3 (also known as PGFS and PTGFS) AKT serine/threonine kinase AMP activated protein kinase angiopoietin activating transcription factor 3 autophagy related calcium/calmodulin-dependent protein kinase kinase 2 cyclic adenosine monophosphate caspase 3 carbonyl reductase 1 C-C motif chemokine ligand cluster of differentiation chorionic somatomammotropin hormone C-X-C motif chemokine ligand cytochrome P450 family 11 subfamily A member 1 diacylglycerol DExD/H-box helicase 58 endothelin 1 early growth response 1 estrogen receptor 1 Fas cell surface death receptor fibroblast growth factor 2 Fos proto-oncogene, AP-1 transcription factor subunit forkhead box P3 GATA-binding protein 3 human chorionic gonadotropin hydroxy-delta-5-steroid dehydrogenase, 3 beta- and steroid delta-isomerase 1 indoleamine 2,3-dioxygenase 1 interferon induced with helicase C domain 1 interferon interleukin inositol 1,4,5-triphosphate interferon-stimulated gene ISG15 ubiquitin-like modifier Janus kinase-signal transducer and activator of transcription Jun proto-oncogene, AP-1 transcription factor subunit low-density lipoprotein receptor

The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00017-0

LGALS LH LHCGR LPA MAP1LC3 MAPK MHC miRNA MKI67 MMP MRP MX1 NO NOS NR4A1 OXTR PAG PGE2 PGF2A PGFM PGH2 PGI2 PIP2 PKC PLA2 PLC PMA PRL PRLR PTGES PTGFR PTGS2 PTX3 RIPK RTP4 SCARB1 SERPINE1 StAR STAT TGFB THBS TIMP TNF VEGF WC1

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galectin luteinizing hormone luteinizing hormone/chorionic gonadotropin receptor lysophosphatidic acid microtubule associated protein 1 light chain 3 alpha mitogen-activated protein kinases major histocompatibility complex microRNA marker of proliferation Ki-67 matrix metallopeptidase maternal recognition of pregnancy MX dynamin-like GTPase nitric oxide nitric oxide synthase nuclear receptor subfamily 4 group A member 1 oxytocin receptor pregnancy-associated glycoproteins prostaglandin E2 prostaglandin F2alpha 13,14-dihydro-15-keto-PGF(2alpha) (also known as prostaglandin F metabolite) prostaglandin H2 prostacyclin phosphatidylinositol 4,5-bisphosphate protein kinase C phospholipase A2 phospholipase C phorbol 12-myristate 13-acetate prolactin prolactin receptor prostaglandin E synthase prostaglandin F receptor prostaglandin-endoperoxide synthase 2 pentraxin 3 receptor interacting serine/threonine kinases receptor transporter protein 4 scavenger receptor class B member 1 serpin family E member 1 steroidogenic acute regulatory protein signal transducer and activator of transcription transforming growth factor beta thrombospondin tissue inhibitors of metallopeptidases tumor necrosis factor vascular endothelial growth factor workshop cluster antigen 1

© 2019 Elsevier Inc. All rights reserved.

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17. LUTEOLYSIS AND THE CORPUS LUTEUM OF PREGNANCY

It’s ephemerality is its most distinguishing and important characteristic … and if there is a basic, common system of regulation of all mammalian CL, the clue to its nature must be in what causes ephemerality. Irving Rothchild [1]

INTRODUCTION The corpus luteum (CL) is a transient endocrine gland that produces the steroid hormone progesterone. Since the CL was first described in detail by Regnier deGraaf in 1672 [2], its transient nature has fascinated many generations of scientists. Identification of its origin and function remained elusive until the turn of the 20th century, when Johannes Sobotta determined that it originated from the cells of the ovarian follicle following ovulation. Shortly after this discovery, Gustav Born, Ludwig Fraenkel, and Vilhelm Magnus [3,4] performed a series of ovariectomy and lutectomy surgeries on rabbits and determined that the function of the CL is to maintain pregnancy. Perhaps, the most fascinating aspect of the biology of the CL is its transience. In the absence of a pregnancy, the CL dies, regressing rapidly, ceasing all progesterone production, and leaving in its place a small, scar-like structure called the corpus albicans. However, in the presence of an embryo, regression does not occur, the CL survives, and progesterone remains high, maintaining the pregnancy. Therefore, a detailed understanding of the processes regulating luteolysis is necessary to both fully comprehend the process of normal female reproduction and to address reproductive problems. The process of luteal regression is complex, involving both cessation of progesterone production and cell death. In the absence of a pregnancy, luteolysis is necessary to allow another ovulation to occur and thus, to give the female another opportunity to become pregnant. Here, the mechanisms governing luteal regression in ruminants will be described in detail, with additional reference to mechanisms in pigs, rodents, nonhuman primates, and humans. The mechanisms for initiation of luteal regression, acquisition of the capacity of the CL to regress in response to prostaglandin F2A (PGF2A), and specific cellular events that may mediate luteal regression will be discussed.

ACQUISITION OF LUTEOLYTIC CAPACITY In many species, the CL regresses in response to PGF2A late in the cycle, but fails to regress in response to the same concentration of PGF2A earlier in the cycle. This acquisition of luteolytic capacity [5] has been observed in a variety of species, and from a practical

standpoint, has presented challenges in development of efficient synchronization protocols for livestock, yet it is a phenomenon that is not fully understood. The bovine CL does not regress in response to a luteolytic injection of PGF2A prior to day 6 of the estrous cycle, but later in the cycle, the same concentration of PGF2A will cause it to regress completely and allow another ovulation to occur [6]. Even a double dose of PGF2A does not result in luteal regression on day 5, and although an extended period of progesterone suppression was observed, these CL recovered progesterone production equivalent to the control animals by day 15 of the estrous cycle [7]. Failure to regress in response to PGF2A has also been documented in other species, including the pig, in which the period prior to acquisition of luteolytic capacity lasts for more than half of the luteal phase [8], rat [9], and marmoset monkey [10], although the timing of acquisition of luteolytic capacity differs. Examining differences between the CL that regress and fail to regress in response to PGF2A provides a useful model for understanding mechanisms by which PGF2A induces luteal regression. Interestingly, in the bovine CL, there is no difference in prostaglandin F receptor (PTGFR) expression or affinity for PGF2A in the CL of differing responsiveness to PGF2A [11]. Further, both prior to and after acquisition of luteolytic capacity, an injection of PGF2A reduced expression of steroidogenic enzymes and abundance of ascorbic acid in the CL [12]. These findings indicate that the early CL possesses the ability to respond to PGF2A, yet they do not regress. Luteolytic capacity may be mediated by the ability of PGF2A to induce PTGS2, a prostaglandin synthesis enzyme, because in day 4 CL, PTGS2 was reduced by PGF2A, but in day 10 CL, PTGS2 was increased by PGF2A [12]. The ability of the CL to synthesize intraluteal prostaglandins is a requirement for structural regression [13]. While the early CL has greater concentrations of both PGF2A and 6-keto PGF1A (the stable metabolite of PGI2) than midcycle or late CL, the ratio of PGI2 to PGF2A is much greater in the early CL than in the fully functional CL [14] indicating that the balance of intraluteal luteotropic versus luteolytic prostaglandins may be an important component of acquisition of luteolytic capacity. Alteration of intracellular calcium homeostasis is a downstream effect of PGF2A in the CL. There was differential regulation of intracellular calcium homeostasis in developing and mature CL [15] and CAMKK2 was expressed in greater abundance by the fully functional CL than by the developing CL and was induced by PGF2A only in the fully functional CL [16]. CAMKK2 activates the signaling molecule AMPK and AMPK itself was also more abundant in day 10 than day 4 CL. In vitro, in day 4 CL, pharmacological AMPK activators were unable to reduce progesterone concentrations, but in day 10 CL, pharmacological activation of AMPK reduced

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progesterone production. PGF2A treatment resulted in AMPK phosphorylation, via CAMKK2, indicating a PGF2A effect on AMPK activation. This phosphorylation led to a decrease in LDLR expression and an increase in ACAT1, which together would make cholesterol less available for steroidogenesis [17]. Overall, these results indicate that as CL acquires luteolytic capacity, AMPK gains the ability to be activated by PGF2A and regulates downstream effects, including progesterone production. Luteal cytokine production, and thus perhaps infiltration of immune cells, in response to PGF2A is also different in the early CL, compared with the fully functional CL. PGF2A induced CCL2, a chemokine that induces tissue infiltration of monocytes, only in mature ovine CL, but failed to induce CCL2 in the CL that had not yet acquired luteolytic capacity [18]. In a recent wholetranscriptome study of the bovine CL on days 4 and 11, before and after a luteolytic injection of PGF2A, functions of mRNA that were differentially expressed on day 11, but not on day 4, after PGF2A injection included functions related to immune regulation and apoptosis. Interestingly, there was a subset of mRNA regulated by PGF2A only on day 4 and a subset regulated only on day 11, with some overlap between the two groups [19]. These data, like the early studies of acquisition of luteolytic capacity, indicate that day 4 CL does respond to PGF2A, but this response does not result in the demise of the CL. Luteal vasculature also responds differently to PGF2A after acquisition of luteolytic capacity. In the CL that regressed in response to PGF2A, there was a rapid increase in blood flow for the first 2 h following injection of PGF2A, followed by a decline in blood flow thereafter, whereas in the CL that failed to regress in response to PGF2A, PGF2A injection did not alter blood flow [20]. The potent vasoconstrictor EDN1 and its receptor were also induced in the midcycle CL in response to PGF2A, but not in the early CL [21,22]. The early CL may have mechanisms that stabilize vasculature against the effects of PGF2A. Similar to the CL that has not yet acquired luteolytic capacity, mature CL may be exposed to PGF2A, but not regress. Atli et al. [23] developed a model for investigating temporal changes in the CL that regress and do not regress in response to subluteolytic pulses of PGF2A, again providing evidence for luteolytic and nonluteolytic actions of PGF2A. In this study, samples were collected from the CL exposed to 0, 2, or 4 intrauterine infusions of PGF2A over 24 h. All the CL that were exposed to PGF2A had differentially expressed early response genes in response to PGF2A, including JUN, FOS, EGR1, NR4A1, and all the CL that experienced four pulses of PGF2A regressed. However, following two pulses of PGF2A, some CL regressed while others did not, and these CL of differing fates had differences in gene

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expression. In the CL that regressed, but not in the CL that failed to regress, there was immune activation (indicated by upregulation of IL1B and CXCL8), increased synthesis of prostaglandins (as indicated by upregulation of PTGS2 and AKR1C3), and decreased vascular stability (as indicated by downregulation of VEGFA) [23], indicating that these factors may be key determinants of whether luteal regression progresses. Similarly, EDN1-treated ewes underwent luteal regression in response to a dose of PGF2A that failed to induce luteal regression in untreated ewes [24]. Interestingly these key pathways are similar to those that are differentially regulated prior to and after acquisition of luteolytic capacity. Overall, for luteal regression to occur, responses to PGF2A must include altered prostaglandin synthesis and metabolism, immune cell chemoattraction and activation, alteration of vasculature, and specific cell signaling events. For a summary of events that occur in response to PGF2A before and after acquisition of luteolytic capacity, see Fig. 1.

INITIATION OF LUTEOLYSIS In most domestic animals, the uterus is required for luteal regression. This was first demonstrated when it was observed that the CL was maintained for a much longer period of time in hysterectomized than in intact guinea pigs [25]. Subsequently, Wiltbank and Casida [26] demonstrated the same phenomenon in the cow and sheep. The work of many researchers collectively demonstrated that in these species, PGF2A is the uterine luteolysin, and the effect is a local one [27–30]. In the ruminant, horse, and pig, amplitude and frequency of pulsatile PGF2A from the uterus reaches the necessary magnitude to induce regression of the CL and loss of progesterone. For complete luteal regression to occur, repeated pulsatile release of PGF2A from the uterus is required. Ginther et al. [31] measured six sequential pulses of PGFM, a PGF2A metabolite that is often measured in place of PGF2A in the blood, during spontaneous regression. In this same study, this research group also demonstrated that one 0.5 mg pulse of PGF2A induces a temporary decline in progesterone, but fails to induce regression, while four identical pulses, each separated from the next by 12 h, induce complete luteal regression [31]. Much less is understood about the mechanism by which the CL of humans and nonhuman primates undergoes regression; however, it is known that the human ovary will continue to cycle normally in the absence of the uterus. Pituitary oxytocin is likely the pacemaker for the pulsatile release of PGF2A from the endometrium in sheep [32]. In ewes, during luteal regression, PGFM and neurophysin 1 and 2 (used as a proxy for oxytocin concentration) pulses measured in the blood occurred

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FIG. 1 Corpora lutea express prostaglandin F receptor (PTGFR) and respond to PGF2A both before and after acquisition of luteolytic capacity. (A) Responses that are induced by PGF2A in the CL both prior to and after acquisition of luteolytic capacity. (B) Effects of PGF2A that differ in the CL before (left box) and after (right box) acquisition of luteolytic capacity. (C) Differences in mature CL that regress or do not regress in response to the same number of pulses of PGF2A (adapted from Atli MO, Bender RW, Mehta V, Bastos MR, Luo W, Vezina CM, et al. Patterns of gene expression in the bovine corpus luteum following repeated intrauterine infusions of low doses of prostaglandin F2alpha. Biol Reprod 2012;86130(4):1–13), showing key pathways that must be activated to induce regression.

(A) (B)

(C)

concurrently [33]. However, in heifers, only half of oxytocin and PGFM pulses occurred concurrently [34]. Further, an oxytocin receptor inhibitor infused into the uterus, at a concentration validated to suppress oxytocin-induced PGF2A release, altered neither cycle length nor plasma progesterone in cattle [35]. Furthermore, oxytocin infusion into the aorta abdominalis that resulted in plasma concentrations of oxytocin equivalent to those observed during luteolysis did not result in PGFM pulses, while a supraphysiological concentration of oxytocin, did induce PGFM in heifers on day 17 of the estrous cycle [34]. Therefore, some controversy remains regarding the widely accepted hypothesis that oxytocin is the initiator of luteal regression in cows.

SPECIFIC LUTEOLYTIC ACTIONS OF PGF2A ON THE DIVERSE CELL TYPES OF THE CL Although discussion of luteolysis is sometimes subdivided into functional (loss of progesterone) and structural (cell death and involution) components, in reality these two processes are intimately linked. However, the rapid decrease in progesterone production begins prior to onset of cell death [36], meaning that death of steroidogenic cells is not the initial cause of the decline in progesterone

production but that PGF2A directly decreases progesterone. Therefore, specific actions of PGF2A on each cell type in the CL, which ultimately result in luteal regression, will be discussed, rather than subdividing these actions into functional and structural actions.

Actions of PGF2A on Steroidogenic Cells In vitro studies of the PGF2A-induced reduction of steroid hormone production have investigated LH responsiveness, lipoprotein utilization, and intracellular signaling events, and have provided an excellent model of ways in which PGF2A induces rapid loss of progesterone production in vivo. Although PGF2A reduces steroidogenic enzyme and LHCGR expression [12,37,38], these events occur after the decline in progesterone production, meaning that this decrease must be mediated by rapid intracellular signaling. Contrary to expectations based on the in vivo observation that one supraphysiological injection of PGF2A results in complete luteal regression when given at the appropriate phase of the estrous cycle in most species, PGF2A fails to kill cultured luteal steroidogenic cells and does not reduce basal production of progesterone in vitro. Although this observation might draw into question the idea that PGF2A is the primary luteolysin, evidence that PGF2A is luteolytic in vivo is quite

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conclusive [27–30]. Measurements of PGF2A and PGFM at the time of luteal regression indicate clearly that PGF2A pulses increase at the time of luteal regression, compared with times earlier in the estrous cycle [39,40]. Furthermore, exogenous PGF2A injection consistently results in complete luteolysis and mice lacking PTGFR have failed parturition, but parturition can be induced by ovariectomy [41]. However, recapitulation of the luteolytic process in vitro has been challenging, because PGF2A does not directly kill luteal steroidogenic or luteal endothelial cells in vitro and careful consideration of cell culture system and concentration of PGF2A treatments must be made in order to see a physiological-like decrease in progesterone induced by PGF2A. Although PGF2A did not reduce basal progesterone production in vitro, it inhibited LH-stimulated progesterone production [42,43]. Similarly, cultured luteal cells from the CL collected either 1 or 4 h after an injection of PGF2A were nonresponsive to LH [44]. This inhibition seems to take place after accumulation of cAMP, as PGF2A also inhibited dibutyryl cAMP-stimulated progesterone production [43]. The mechanism of action of the observed inhibition is via activation of PKC. PGF2A signals via the PTGFR, a G-protein-coupled receptor, which activates PLC. Activated PLC converts membrane-bound PIP2 to IP3 and DAG. Intracellular calcium concentrations increase in response to IP3, while DAG activates PKC [45]. Activation of PKC leads to activation of MAPK signaling, including MAPK1, MAPK8, and MAPK11 [46–49]. Ultimately, this leads to alteration of transcription factors such as FOS and JUN [50] and changes in gene expression in the regressing CL. Pharmacological activation of PKC by PMA resulted in reduction of basal progesterone production by large luteal cells (LLC) and LH-stimulated progesterone by small luteal cells (SLC), downstream of cAMP accumulation [51] and treatment of LLC with PGF2A increased free intracellular calcium [52]. It has been hypothesized that the increase in intracellular calcium ions mediates PGF2Ainduced cell death in LLC, because the calcium ionophore A23187 resulted in death of LLC, but not SLC [53]. However, infusion of PMA, which activates PKC, into the ovarian artery transiently reduced progesterone production, but did not result in formation of apoptotic oligonucleosomes or luteolysis, while infused PGF2A reduced progesterone production and did induce apoptosis and luteolysis [54]. This indicates that activation of PKC may be a mechanism by which PGF2A reduces progesterone production, but it is not the luteolytic mechanism per se. Further, given the finding that PGF2A fails to kill luteal cells in vitro, despite inducing intracellular calcium [52] indicates that there are additional signaling mechanisms required for intracellular calcium to result in cell death. An important component of the loss of progesterone production in the rodent, but not in most domestic

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large animals or humans, is induction of the progesterone metabolizing enzyme AKR1C1 [55], which in the rodent explains the rapid loss of functional progesterone signaling; progesterone is rapidly converted to its inactive metabolite, 20-alpha-hydroxy-progesterone. Another mechanism of action of PGF2A is to inhibit the ability of luteal cells to utilize lipoproteins for steroid hormone biosynthesis. Low-density lipoprotein treatment induced progesterone production in vitro, but pretreatment with PGF2A inhibited this increase, without inhibiting lipoprotein uptake [44,56]. This is supported by the finding that lipid droplets accumulate in the cytoplasm of luteal cells during regression [57]. The mechanism by which PGF2A inhibits lipoprotein utilization for steroidogenesis occurs after cholesterol transport to the mitochondria, but prior to cholesterol side-chain cleavage by CYP11A1 [58]. Interestingly, ATF3, which increases in abundance as early as 1 h after induction of luteal regression, also inhibited progesterone production prior to cholesterol side-chain cleavage, but without altering expression of the StAR protein [59]. ATF3 may be an early mediator of the decrease in progesterone production observed in luteal regression. Cholesterol efflux is also a component of the decrease in progesterone production observed during luteolysis; ABCA1, a regulator of cholesterol efflux, increased during luteal regression [60]. Mechanisms by which PGF2A alters cellular functions are depicted in Fig. 2. Ovine SLC express the LHCGR in greater abundance and the PTGFR in lesser abundance than LLC. Greater expression of the LHCGR makes the SLC even more LH responsive than LLC, although LLC produce greater basal concentrations of progesterone [61]. Given that LLC express the PTGFR even more abundantly than SLC, LLC are likely to mediate the PGF2A-induced decrease in progesterone production. Niswender et al. [13] suggested that LLC indeed respond to PGF2A and that the decrease in LH responsiveness of SLC in the presence of PGF2A is regulated by LLC. At high concentrations, which are used commonly in culture systems, PGF2A induces progesterone production and protects cultured luteal cells from apoptosis, a counterintuitive finding [62], perhaps by binding to the prostaglandin E receptor. Concentrations of PGF2A found in the uteroovarian venous plasma during luteolysis in the sheep reach 22 ng/mL at the peak of the highest concentration PGF2A pulse [63], so this likely reflects the concentration of PGF2A that reaches the CL. A common concentration of PGF2A used in in vitro studies is 1 μM (354 ng/mL). PGE2 is a known luteotrophin and it has been demonstrated that PGF2A can bind to the PGE2 receptor [64] and will bind to the PGE2 receptor at concentrations much lower than 1 μM [65]. Furthermore, a pharmacological concentration of PGF2A induced progesterone production in SLC but suppressed LH

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FIG. 2

Downstream effects of PGF2A signaling in luteal cells. PGF2A binds to the PTGFR, a G-protein-coupled receptor, which leads to activation of PLC and PLA2. Activation of PLC induces release of IP3 and DAG, which increase intracellular calcium concentrations and activate PKC, respectively. Activation of PKC induces components of MAPK signaling, including MAPK1, MAPK11, and MAPK8, which leads to upregulation of the nuclear transcription factors FOS, JUN, NR4A1, EGR1, and ATF3. FOS, JUN, NR4A1, and EGR1 induce transcription of genes associated with apoptosis and inflammation, and ATF3 reduces LH-stimulated progesterone production. MAPK signaling also leads to upregulation of CXCL8, a cytokine that induces neutrophil activation and migration. The increased intracellular concentrations of calcium induced by IP3 indirectly lead to the activation of CAMKK2, which phosphorylates AMPK. AMPK signaling leads to decreased concentrations of LDLR and SCARB1 and increased concentrations of ACAT1 and ABCA1, which together lead to reduced availability of cholesterol precursors for steroidogenesis. Under the influence of PGF2A, there is also reduced transport of cholesterol into the mitochondria for cleavage by CYP11A1. Activation of PLA2 leads to release of arachidonic acid from the cellular membrane, to be used as a precursor for prostaglandin biosynthesis. This, coupled with an increase in PTGS2, leads to increased availability of PGH2. PGH2 may be converted to PGE2 by PTGES or to PGF2A by AKR1C3. CBR1 converts PGE2 to PGF2A. Increases in PTGES, AKR1C3, and CBR1 during luteal regression lead to a PGF2A autoamplification loop.

responsiveness in LLC [66]. As discussed previously, LLC have the PTGFR while SLC lack the PTGFR or express it at low concentrations [61], indicating that perhaps the mechanism by which PGF2A induces progesterone at supraphysiological concentrations is not mediated by the PTGFR.

Actions of PGF2A on Endothelial Cells The CL has a rich blood supply. Substantial angioregression occurs during luteolysis [67], with changes in luteal blood flow taking place very early during luteal regression [20]. Luteal endothelial cells are the first to undergo apoptosis [68]. These and other data demonstrate the importance of vascular changes during luteal regression. Although there have been conflicting reports about the presence of PTGFR on luteal endothelial cells [69,70], recent reports have convincingly used immunohistochemistry to localize PTGFR expression on both luteal steroidogenic and luteal endothelial cells [71] and have demonstrated PTGFR mRNA expression in

cultured endothelial cells, both in the presence and absence of cytokine treatment [72]. The previously reported discrepancies may be due to variation in the specific type of endothelial cell, among the five types identified [73], used in the various reports and laboratories. These data indicate that, most likely, luteal endothelial cells (or at least, some proportion of the luteal endothelial cell population) have the ability to respond directly to PGF2A. Angiotensin 2 and EDN1 are vasoactive peptides, both of which are induced in luteal regression [74,75]. Endothelial cells themselves are rich sources of EDN1, while steroidogenic cells are sources of both peptides [75]. Both peptides are potent vasoconstrictors and are proposed to contribute to the loss of bloodflow seen during luteal regression. EDN1 is expressed more abundantly in the CL at the end of the estrous cycle (day 18) and during induced luteal regression than it is in the fully functional CL. in vitro treatment with PGF2A induced EDN1 in luteal endothelial cells [74]. Similarly, angiotensin 2, as well as the enzyme responsible for cleavage of its biologically inactive precursor, are induced during luteal

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regression [75]. EDN1 reduced both basal and LH-stimulated progesterone production by luteal steroidogenic cells in a dose-dependent manner [76], whereas angiotensin 2 reduced LH-stimulated, but not basal progesterone production [77]. Moreover, injection of the CL with an endothelin receptor inhibitor delayed the onset of luteal regression in response to PGF2A and treatment with EDN1 itself resulted in reduced progesterone production and increased luteal sensitivity to a subluteolytic dose of PGF2A [24]. Overall, these data indicate an important role for vasoconstrictive peptides in luteal regression. NOS and its production of NO also seem to be important components of the luteolytic cascade. NO is a vasodilator and its proposed role is to modulate the transient increase in bloodflow in the CL during early luteolysis. Treatment with a NOS inhibitor abrogated the transient increase in bloodflow seen in the periphery of the CL during luteal regression, delayed the onset of decrease in plasma progesterone and the onset of decreased luteal size, but did not prevent luteal regression [78]. Perhaps, NO synthesis hastens the luteolytic process, but is not obligatory for the progression of luteal regression. Supporting this idea, in a study that compared the luteolytic actions of PGF2A analogs dinoprost and cloprostenol, both induced luteal regression, but only dinoprost induced the rapid increase in peripheral blood flow previously reported [79], indicating that this increase is not obligatory for the completion of luteal regression. Various angiogenesis-related factors are also regulated during luteolysis. Angiopoietins are vascular growth factors, the functions of which are regulated primarily by the ratio of ANGPT1 to ANGPT2, with greater concentrations of ANGPT1 leading to blood vessel formation or stabilization and greater concentrations of ANGPT2 leading to blood vessel destabilization and regression in the absence of vascular endothelial growth factor (VEGF). The ratio of ANGPT2 to ANGPT1, an indication of loss of stability of blood vessels, is greater in spontaneous regression (CL staged later than day 18) and in induced luteal regression [80] than in the fully functional CL. VEGF, a major angiogenic factor in the formation of the CL, is also reduced during luteal regression [81]. Contrary to expectation, another proangiogenic factor, (FGF2), is induced during luteal regression [81,82]. However, FGF2 is induced to a much greater extent by PGF2A in the CL that are refractory to PGF2A than in the CL that regress in response to PGF2A. Further, the proangiogenic actions of FGF2 are likely somewhat inhibited during luteal regression, but not inhibited following PGF2A exposure in the developing CL, because two specific inhibitors of FGF2 action, PTX3 and THBS were also induced during luteal regression [82]. These factors may serve to sequester FGF2 and counter its proangiogenic actions during luteal regression.

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THBS has important proapoptotic functions during luteal regression, intimately linked to those of TGFB. TGFB1 caused reduction of angiogenic potential [83] and increased cell death in luteal endothelial cells in vitro [83,84]. TGFB1 and THBS1 were regulated by PGF2A in vivo during luteal regression, with THBS1 increasing to a much greater extent and more stably after an injection of PGF2A in the CL that regress in response to PGF2A than in early, refractory CL. THBS2, but not THBS1, was regulated by a PGF2A analog (cloprostenol) in vitro in luteal endothelial cells [82]. Interestingly, although PGF2A seemed to induce thrombospondins without inducing cell death in vitro, exogenous THBS1 treatment reduced cell viability in cultured luteal endothelial cells [84], with the highest concentration of THBS1 decreasing viability over 80%. TGFB1 was activated by THBS1 and induced transcription of SERPINE1, a profibrotic factor that has been previously reported to be associated with luteolysis [85]. However, a lesser proportion of cells died in response to TGFB1 treatment than in response to THBS1 treatment [84], indicating that THBS1 likely has other important downstream targets to induce cell death, rather than simply inducing activation of TGFB1. In addition, TGFB1 increased expression of collagen and matrix metalloproteinases in luteal fibroblasts [86], which indicates that it may contribute to the tissue remodeling which takes place during structural regression

SPECIFIC ACTIONS OF IMMUNE CELLS AND CYTOKINES IN LUTEOLYSIS Lobel and Levy were the first to report the presence of immune cells in the CL, in 1968 [87]. Since this initial report, it has been demonstrated that total immune cell number in the CL increases modestly during luteal regression [88], but not late in the estrous cycle [89] and that there is alteration in production of cytokines and their receptors in the CL during luteal regression. Bauer et al. [90] used immunohistochemistry and colocalized the marker of cell-cycle progression, MKI67, with various immune cell markers in vivo. They reported that although there was an increase in proliferation of immune cells during luteal regression in vivo, this increase was due to proliferating CD14+ cells (likely macrophages) in late regression [90], indicating that both infiltration and proliferation are sources of luteal immune cells. While PGF2A induces luteal regression in vivo, it kills neither luteal steroidogenic cells nor luteal endothelial cells in vitro. Cytokines, unlike PGF2A, induce cell death in cultured luteal cells and because they are produced at greater abundance in luteal regression than during the estrous cycle, may be key regulators of luteolysis.

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Immune cells The CL is home to a diverse population of immune cells, including T lymphocytes, macrophages, neutrophils, and eosinophils. Some of these become luteal resident immune cells, while others are more transient and may only be in the CL for a short time prior to their death. In general, luteal immune cell abundance increases during luteal regression. Considerable in vitro characterization of T cell-luteal cell interactions has been carried out, while less is known about luteal cell-macrophage or luteal cell-neutrophil interactions. Reports of PTGFR expression on peripheral immune cells are conflicting [69,91]. While luteal resident immune cells may be programmed by PGF2A during luteal regression, there is likely a very important indirect effect of the luteal microenvironment (luteal steroidogenic and endothelial cells and their products) upon programming of luteal resident immune cells. Total T cells (CD5+) and CD8+ T cells were greater in abundance in the CL during spontaneous regression (days 19–21) than in the CL of the early, mid-, or late estrous cycle in cattle. Interestingly, this increase in CD5+ and CD8+ cells was not recapitulated in induced regression [88]. Regardless of status (functional or regressing), the CL had a greater proportion of CD8+ and a smaller proportion of CD4+ T cells than the peripheral blood [92]. In rhesus monkeys, CD3+ T cells increased during regression, but not until 3–4 days after the initial decline in progesterone [93]. However, T cells are present in the CL throughout its lifespan and have been demonstrated to induce progesterone production [94], likely through production of luteotropic cytokines, such as IL4 and IL10 [95], indicating that in some contexts they are supportive of luteal function. However, luteal cells are potent stimulators of bloodderived autologous T cell proliferation in vitro [96] and luteal cells derived from regressing CL stimulated greater proliferation than luteal cells derived from midcycle CL, regardless of whether the T cells were collected prior to or after injection of PGF2A [97], demonstrating that this is a luteal cell-specific, not T cell-specific, effect of PGF2A. MHC1 molecules, as well as the costimulatory molecules, CD80 and CD86, appear to mediate at least some of the luteal cell-stimulated T cell proliferation, because blocking these molecules with antibodies attenuated luteal cellinduced T cell proliferation [98,99]. However, the cells that respond to the greatest extent to luteal cells are the nonMHC-restricted gamma delta subset of T cells [98]. It is unknown what molecule(s) are used by luteal cells to activate gamma delta T cells. In ruminants, subpopulations of gamma delta T cells can be distinguished by expression of WC1. Gene expression arrays have indicated that WC1+ cells may be of a more pro-inflammatory types, whereas WC1
cells are more antiinflammatory [100]. In coculture

experiments, steroidogenic cells derived from a fully functional CL preferentially induced proliferation of WC1
gamma delta T cells and steroidogenic cells derived from a regressing CL preferentially induced proliferation of WC1+ gamma delta T cells. In addition, midcycle but not regressing luteal cells, induced gamma delta T cells to produce IL10 and express GATA3, which suppress T cell proliferation and pro-inflammatory cytokine production [101]. The luteal microenvironment in a functional CL may program gamma delta T cells to such that they facilitate homeostasis within the tissue, rather than promote an inflammatory response. Programming of T cells within the luteal microenvironment may be a key component of luteal resident T cell function. For example, regression increased the proportion of CD8αα+ total T cells. CD8αα+ T cells are resolving-type T cells and may increase during luteal regression in response to an inflammatory-type response that may occur immediately after the onset of regression. Importantly, there was a greater than fivefold increase in FOXP3+ (T regulatory) cells among luteal resident T cells derived from a fully functional CL compared with luteal resident T cells derived from a regressing CL or peripheral T cells from either functional status [92], indicating that resident T cells may be more regulatory in a fully functional CL than in a regressing CL and that there is likely a role for the luteal microenvironment in programming these cells to a more regulatory phenotype. Progesterone has functional effects on T cells [97,102–104]. While T cells do not express nuclear progesterone receptors, they do express membrane progesterone receptors [104]. The high endogenous concentration of progesterone in the CL may program the resident T cells. There is macrophage infiltration into the CL in the middle to late estrous cycle [88,89]. Reports of macrophage numbers changing in regressing CL are conflicting [88,105]. Cytokine production [106] and withdrawal of steroid hormones [107] may be important components of macrophage infiltration. During late regression, there were more proliferating CD14+ macrophages than during the estrous cycle or early in regression, indicating that an important source of the macrophages seen in late regression may be cell proliferation [90]. Macrophages may be important sources of cytokines involved in luteal regression; for example, these cells are sources of tumor necrosis factor (TNF), a cytokine that has a key role in the progression of luteal regression. Recent evidence from the primate, however, indicates that macrophages may be more important in the structural than functional changes associated with luteolysis; macrophage infiltration does not occur in the late luteal phase, rather it occurs at menses [108] or after progesterone has been low for several days [93]. Interestingly, macrophages seemed to be associated with areas of cell death and apoptosis and were absent in portions of the tissue that were still healthy,

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in tissue sections taken from regressing CL [108], indicating that perhaps the role of macrophages is to clean up dying or dead cells in the CL as regression progresses. CCL2 is a chemokine that is expressed by luteal endothelial cells or, in some species, by luteal steroidogenic cells [89,108,109] increasing abundance as the estrous cycle progresses. CCL2 is associated with attraction and infiltration of immune cells, particularly monocytes or macrophages. Its abundance is increased during luteal regression in sheep [109], rhesus monkeys [110] and women [108,111], and in induced [112] and spontaneous [113] regression in cattle. In the bovine estrous cycle, the increase in expression of CCL2 precedes the onset of spontaneous regression; pronounced increase in CCL2 transcript was observed comparing days 6–12 of the estrous cycle, with no further change from days 12 to 18 [89], indicating that there is infiltration of immune cells regulated by CCL2 prior to luteal regression. Furthermore, in the bovine CL, CCL2 does not seem to be regulated by PGF2A directly, but is induced by the cytokines TNF and IFNG [114]. In women, the abundance of CCL2 seen in the late phase CL was reduced by exogenous administration of hCG for several days prior to hysterectomy and sample collection [108], indicating that perhaps an important component of luteal rescue is prevention of the induction of CCL2. There is an acute infiltration of neutrophils [115] and eosinophils [116] into the CL during early luteolysis. Their function is not entirely clear, although it has been demonstrated that neutrophils do not reduce basal or LH-stimulated progesterone production [117], and in fact, IFNT-treated neutrophils increase progesterone production [118]. Perhaps, the role of these cells is to mediate early inflammatory events during luteal regression. Although much work has been done to determine the phenotype and function of luteal resident immune cells, it has been difficult to elucidate their role in the regulation of luteal steroidogenic cell function, progesterone production, and response to endocrine and paracrine factors. Blood-derived lymphocytes and monocytes stimulated progesterone production from granulosal cells [119]. However, PBMC that were activated with concanavalin A abrogated LH-responsiveness of cultured luteal cells, but did not alter basal progesterone production [117].

Cytokines Bagavandoss et al. [120] were the first to investigate the role of immune cell-derived cytokines in luteal function when they reported the presence of TNF in leporine corpora lutea and greater ability of regressing CL to produce TNF in response to stimulation with LPS than the early CL or the CL of pregnancy [120,121]. Since those first reports, an abundance of evidence for cytokine

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involvement in luteal regression has accumulated, including in recent gene profiling studies [19,112]. Cytokine transcripts are differentially expressed during induced luteal regression. TNF, as well as IFNG and IL1B transcript abundance increased from 0 to 2 h after a luteolytic injection of PGF2A and remained elevated for 12 h (IFNG) to 64 h (TNF and IL1B) [116]. TNF, TNF receptor, and IFNG also increased in abundance during spontaneous regression (days 19–21) in the cow, as compared with expression in the CL from earlier in the estrous cycle [122,123]. TNF is also increased in the CL of women during menses, but this increase was prevented by administration of hCG [108]. In a study that used immunohistochemistry to localize TNF and its receptor in the CL, expression of both was demonstrated in large and small steroidogenic cells, as well as immune cells and endothelial cells. Although there was expression of TNF and its receptor in these cell types throughout the estrous cycle, the regressed CL (days 19–21) had even more abundant expression [124]. Perhaps, most compellingly, Henkes et al. [125] demonstrated that pseudopregnant TNF receptor knockout mice failed to undergo PGF2A-induced luteal regression. Similarly, mice treated with a TNF-neutralizing antibody and mice lacking acid sphingomyelinase, a key enzyme in the TNF-induced apoptotic pathway, also did not undergo luteolysis [125]. While it is well established that cytokine production and regulation is a key component of luteolysis, the mechanism by which cytokines may affect luteal function are diverse. One function of intraluteal cytokines may be to induce intraluteal prostaglandin production, which is known to be necessary for structural regression [13]. TNF, IL1B, and IFNG induced prostaglandin production and inhibited LH-stimulated progesterone production by cultured luteal cells, with TNF and IFNG having a particularly potent effect [126–129]. Recently, a lesser-known cytokine, chemerin, has also been demonstrated to be a potential regulator of luteal progesterone production. In incubated luteal tissue, chemerin inhibited hCGinduced and basal progesterone and a blocking antibody directed against the chemerin receptor rescued this effect. Importantly, in vivo treatment with this same blocking antibody rescued the decrease in progesterone and the increase in CASP3 seen with exogenous PGF2A treatment [130], indicating that preventing chemerin signaling may in part block the progression of luteal regression. IFNG, particularly in combination with TNF, was cytotoxic to luteal steroidogenic cells in culture [127,128] and this effect was not inhibited by inhibitors of NO production, prostaglandin production, or phospholipase A2 [131]. However, IFNG and TNF-induced cell death was somewhat rescued by treatment with a free radical scavenger and by treatment with another cytokine, interferon alpha (IFNA) [131], indicating that the mechanism of action of these cytokines may have

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something to do with the accumulation of free radicals and that there may be crosstalk among cytokines as the CL regresses or is rescued. The cytokine-induced cell death seen in these studies was due to apoptosis [131] mediated by the FAS-FAS ligand system [132]. Rapid upregulation of IDO by IFNG mediated IFNG-induced apoptosis by depleting cellular tryptophan [133]. Apoptosis has been observed in luteal regression [36] and FAS mRNA was elevated in regressing CL [132]. Together, these data indicate that cytokines may be key regulators of cell death and loss of progesterone production during luteal regression, particularly given the finding that PGF2A fails to kill luteal steroidogenic cells and luteal endothelial cells at any concentration used in vitro. Luteal endothelial cells are the first cells to undergo apoptosis during luteal regression [68] and loss of luteal vasculature may be an important component of luteal regression, as discussed previously. Although TNF treatment alone was inadequate to kill luteal steroidogenic cells and combination with IFNG was required for death of this cell type, TNF alone or TNF in combination with IFNG, induced apoptosis in luteal endothelial cells [134]. Increases in TNF and IFNG may mediate early changes in vascular stability observed during luteal regression. Matrix remodeling is an important component of luteal regression that may also be modulated by cytokines. MMP was induced during luteolysis, while their regulators, TIMP, were either unchanged or reduced in abundance [135]. IFNG reduced the abundance of TIMP1 and TIMP2, and IFNG and PGF2A synergistically increased MMP1 and MMP9 transcript abundance [136]. These data indicate a possible role for cytokines in matrix remodeling during luteal regression. Although much attention has been focused on the actions of TNF and IFNG in luteal regression, and it is evident that they are important players in the inflammatory events that occur during luteal regression, there are other cytokines and chemokines that have come to the forefront, perhaps most notably CXCL8. CXCL8 increased in abundance during induced [115,117] or spontaneous [123] luteal regression in the bovine CL, and was induced by PGF2A in luteal steroidogenic cells and luteal endothelial cells in vitro [115,117]. CXCL8 was also produced by macrophage/neutrophil populations isolated from the CL of rhesus monkeys [110]. Furthermore, in luteinized human granulosal cells, either PGF2A treatment or withdrawal of luteotropin induced CXCL8 [137]. CXCL8 potently induces neutrophil migration [115,117] and neutrophil infiltration is observed as early as 5 min after PGF2A administration. While increases in CXCL8 were not observed until 30–60 min after administration of PGF2A, CXCL8 could have an important role in later neutrophil infiltration. A variety of cytokines and chemokines are increased during luteal regression. Given that PGF2A does not

directly kill luteal cells in culture, it is likely that the PGF2A-induced increase in the abundance of these cytokines, particularly by resident immune cells, is a key component of the cell death observed during luteal regression. Furthermore, given the observations that cytokines seem to induce prostaglandin production and reduce progesterone production by cultured luteal cells, they are likely involved in the prostaglandin autoamplification loop.

ADDITIONAL MOLECULAR MEDIATORS OF LUTEOLYSIS Steroid hormone biosynthesis involves generation of high intracellular concentrations of free radicals. Dysregulation of the antioxidant enzymes and lipids that reduce these radicals may be involved in apoptosis during luteal regression. In the rodent and the cow, enzymes associated with free radical elimination were downregulated during luteal regression [138,139] and the antioxidant ascorbic acid is depleted from the CL by PGF2A [12,140]. Mice deficient in superoxide dismutase have an increased number of apoptotic luteal cells and decreased plasma progesterone [141]. NO is a potent vasodilator, but can also be a source of free radicals. LPA, which is a phospholipid that is produced locally by the CL, protected against NO- and cytokine-induced decreases in progesterone production and apoptosis markers, increased basal progesterone production, and augmented IFNT stimulation of interferon-stimulated genes (ISGs) in cultured luteal cells [142,143]. These data indicate that LPA may have a luteotropic (progesterone stimulatory) and luteoprotective (prosurvival) role in luteal function. However, finding that although LPA receptors increased in pregnancy, luteal LPA concentration decreased in pregnancy and was at its highest concentration prior to or during luteal regression [142] makes these findings hard to interpret in the context of the normal cycle. Galectins, which characteristically bind to betagalactoside sugars, have recently emerged as regulators of luteal steroidogenic cell death and survival. LGALS3 is induced in spontaneously regressing CL in vivo and by PGF2A in vitro. Treatment of cultured luteal steroidogenic cells with LGALS3 resulted in cleavage of CASP3, a measure of apoptosis, and reduced cell viability [144]. In contrast, it seems that LGALS1 has a luteotrophic effect. It improved cell survival, induced VEGF receptor expression, and was expressed abundantly in the CL until late in the cycle, but dropped off abruptly in regressed CL [144]. Sialic acid binds to LGALS1 recognition sites and blocks LGALS1 binding, a process referred to as alpha 2,6-sialylation. Alpha 2,6-sialic acid is in greater concentration in the late cycle luteal cells (classified as days

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15–17) than in midcycle luteal cells (classified as days 8–12) and, while LGALS1 improved the viability of midcycle luteal cells, it did not improve the viability of the late cycle luteal cells. However, after treatment with neuraminidase, which removes sialic acid from galectin binding sites, LGALS1 improved viability of late luteal cells in a similar way to midcycle luteal cells [145], indicating that removal of sialic acid from LGALS1 binding sites allowed LGALS1 to bind and exert its luteoprotective effect. The balance between LGALS1 and LGALS3, as well as concentration of sialic acid and the protein responsible for sialylation, are likely regulators of luteal cell survival and death.

CELL LOSS DURING LUTEOLYSIS Luteolysis involves extensive cell loss and death. The first type of cell death identified as a component of luteolysis was apoptosis [36]. The FAS-FAS ligand system is the pathway by which cytokine-induced cell death occurs [132] and both the extrinsic and the intrinsic pathways of apoptosis are induced during luteolysis, with mRNA encoding components of the extrinsic pathway elevated earlier than mRNA encoding components of the intrinsic pathway [146]. However, apoptosis is not the only mechanism of cell death during luteolysis. Typically, necrosis is associated with death of damaged tissue, but necroptosis is an orderly, programmed form of necrosis that is not necessarily associated with acute tissue damage. The markers of necroptosis, RIPK1 and 3, were increased in spontaneous regression, as well as 4 h after injection of PGF2A. RIPK1 was induced by treatment with TNF and IFNG, but not by either cytokine alone, and a RIPK1 inhibitor partially protected cultured luteal steroidogenic cells from TNF/IFNG-induced cell death [147]. Another mechanism that may be activated during luteolysis is autophagy, a stress response wherein cells recycle unneeded organelles that can lead to cell breakdown and death. The autophagy markers MAP1LC3A, MAP1LC3B, ATG3, and ATG7, and autophagic activity, were elevated in regressing CL as compared with the fully functional CL [148]. In the rodent CL, markers of autophagy and cleaved CASP3 were elevated at the end of pseudopregnancy [149]. in vitro treatment with PGF2A directly induced the lipid-modified form of MAP1LC3B, and an increase in autophagic cells, via the MAPK signaling pathway [150]. Furthermore, treatment with a pharmacological autophagy promoter increased luteal cell death while a pharmacological autophagy inhibitor reduced cell death [149]. Another novel mechanism that has been suggested is loss of cells into the lymphatic fluid drainage from the ovary. Live, HSD3B1 positive, lipid droplet-containing cells were identified in the lymphatic fluid drainage from the regressing CL

[151]. These authors propose that this mechanism of loss may be important to ensure that the CL disappears completely, leaving only a corpus albicans. Overall, these relatively new findings demonstrate that apoptosis alone does not account for the cell loss during luteal regression and that other mechanisms are also components.

LUTEAL RESCUE Maternal recognition of pregnancy (MRP) is the process by which the reproductive system of a mother recognizes that a conceptus is present and makes the adjustments necessary to maintain the pregnancy. An essential event during MRP is rescue of the CL and thus, continuation of luteal progesterone production, which maintains pregnancy. Although the ultimate outcome— that is to say, luteal rescue—of conceptus-derived MRP signals is shared, there is a tremendous diversity of MRP signals in mammals. This review will discuss mechanisms of luteal rescue in ruminants and humans in depth and will briefly discuss what is known about other species, with references to recent reviews.

Luteal Rescue in the Ruminant In the ruminant, the primary MRP signal is a trophoblast-derived interferon, IFNT. IFNT production by the trophectoderm of the conceptus begins around days 14–15 in cattle [152] and slightly earlier in sheep. Embryo transfer pregnancies in cattle survive when transfer takes place prior to or on day 16 of pregnancy, but are lost when transfer takes place on or after day 17, demonstrating the critical timing of onset of IFNT production [153]. A similar mechanism is seen in the sheep [154]. The specificity of timing of the MRP signal results from the necessity of rescuing the CL from luteolytic pulses of uterine PGF2A in these species. The actions of conceptus-derived hormones on the uterus, and perhaps also the CL, are necessary for luteal rescue. Antiluteolytic Actions of IFNT on the Uterus Intrauterine infusion of conceptus secretory products or IFNT prevents luteolysis in sheep [155,156] and cows [157]. In early studies of the effects of conceptus secretory proteins either enriched for, or depleted of, IFNT, IFNT was the only conceptus-derived substance that prevented oxytocin- or estradiol-induced luteolysis. It is evident that in both the ewe and the cow, exogenous conceptus secretory products or IFNT attenuated plasma PGFM and very likely uterine PGF2A release in response to an exogenous stimulus such as oxytocin or estradiol [155,156,158]. However, these findings must be considered in light of research regarding PGF2A content and release in cyclic

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and pregnant uteri, without exogenous stimulation. Lewis et al. [159] demonstrated that PGF2A content was greater in the ovine endometrium of pregnancy than the estrous cycle, while there was no effect of pregnancy on uterine venous plasma PGF2A on days 11–16. In heifers, PGF2A concentration modestly reduced uterine venous plasma during pregnancy compared with the cycle, although concentration in the ovarian artery was not different [160]. However, PGF2A concentration in uterine flushings was greater in pregnant than cyclic cattle [161]. Further, there is no evidence for conceptus regulation of prostaglandin synthase expression [162]. Despite the lack of difference in PGF2A concentration during the estrous cycle and pregnancy in ewes, there seems to be a difference in pulsatile release of PGF2A, as pulses were fewer and interpulse intervals were longer in pregnant ewes than in cyclic ewes [163]. Given that multiple pulses of PGF2A are required for luteal regression [31], this alteration in release of PGF2A likely explains the antiluteolytic effect of the conceptus in early pregnancy. Interferon response elements control expression of ESR1, which in turn regulates expression of OXTR and pulsatile release of PGF2A from the uterus [164,165]. However, the mechanism of IFNT-mediated alteration of PGF2A may be somewhat different in the cow. While there was lesser plasma PGFM in response to oxytocin injection in pregnant than in cyclic or bred, nonpregnant cows, detection of OXTR in the endometrium of heifers was limited, OXTR protein was detectable in only 2/14 open cows. In addition, there was no effect of pregnancy on ESR1 expression in cows [166]. This indicates that if oxytocin signaling is involved in repression of luteolysis in cows, the mechanism may differ from that in the sheep. In addition, a recent study demonstrated a much greater PGE2:PGF2A ratio in the uteroovarian venous plasma and ovarian arterial plasma of pregnant than cyclic ewes, indicating greater delivery and perhaps preferential transport of PGE2 to the CL of pregnancy [167], albeit this was on day 16, which was likely during luteal regression in the nonpregnant animals. IFNT, via the MAPK signaling pathway, inhibited the function of the prostaglandin transporter protein, which mediates the release of prostaglandins from cells, in endometrial luminal epithelial cells [168]. Overall, these studies indicate that altered prostaglandin release and transport may be an important component of the antiluteolytic mechanism mediated by IFNT. Direct Actions of IFNT on the CL That May Render It Less Sensitive to PGF2A Uterine release of PGF is not ablated during early pregnancy and may not even be reduced, indicating that the CL of pregnancy is exposed to a concentration of PGF2A equal to that capable of inducing luteolysis in the estrous cycle. The ovine [169,170] and bovine [171] CL of

pregnancy are more resistant to the luteolytic actions of PGF2A than the CL of the cycle. In addition to its actions on the uterine endometrium and uterine glands, IFNT also escapes the uterine environment [172] and may have important actions on blood cells or distant tissues, including the CL. Interferon tau is very lowly abundant in the bloodstream and had not been successfully measured in the periphery until recently, so expression of ISG mRNA or protein compared with a control has been used as a measure of interferon exposure in peripheral tissues and immune cells. ISGs, including ISG15, MX1 [173], DDX58, IFIH1 [172], and RTP4 [174] are detectable in the CL as early as day 14 of pregnancy in the sheep. ISGs are also detectable in the CL following infusion of IFNT into an open uterus [175], which indicates that the upregulation of ISGs in the CL during pregnancy is very likely an IFNT-mediated event. Although it has been demonstrated previously that intrauterine infusion of IFNT extended luteal lifespan, the observation that uterine vein or jugular vein infusion of IFNT protected the CL from exogenous luteolytic PGF2A indicates a potential endocrine role for IFNT [176,177]. This is in agreement with evidence that the ovine CL of pregnancy is more resistant to the actions of PGF2A than the CL of the cycle and that this resistance is transient and is lost between days 19 and 26 of pregnancy [169,170], perhaps due to the decline in IFNT production. IFNT directly alters gene expression in luteal steroidogenic cells. Treatment of cultured luteal steroidogenic cells with interferon tau did not alter progesterone production [118], but did increase expression of ISGs in several studies [118,173,178]. In the CL of pregnancy or after infusion of IFNT, ISG15 expression is abundant in LLC and somewhat lesser in SLC [177,179]. This observation indicates that ISG expression in the CL of pregnancy may not be simply due to infiltration of immune cells that have been exposed to IFNT in the periphery, because there is induction of ISGs in luteal cells upon direct IFNT exposure. IFNA is a type I interferon, like IFNT, and signals through the same receptor. Interestingly, both IFNT and IFNA extended cycle length in ewes when infused into the uterine lumen, but ewes that received IFNT seemed to have longer extension of their cycles than ewes that received IFNA [180]. IFNA protected cultured luteal cells from IFNG/TNF-induced cell death [131], suppressed TNF- and TNF and IFNG-induced PGF2A production in cultured luteal cells [181], and reduced IFNG-induced upregulation of MHCII molecules in cultured luteal steroidogenic cells [182]. These data indicate that perhaps type I interferons protect luteal cells from the cytotoxic effects of cytokines during the MRP period. Luteal endothelial cells also seem to be affected by IFNT. Cultured luteal endothelial cells respond to IFNT treatment with increased proliferation and upregulation

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of ISG [183]. Interestingly, IFNT also downregulated four transcripts that are induced during luteolysis, including TGFB1, THBS1, SERPINE1, and EDN1. In luteal tissue slices, THBS1 and SERPINE1 were induced by PGF2A, but in the presence of IFNT, this increase was abrogated, suggesting a possible protective effect of IFNT against some actions of PGF2A [183]. By day 40 of pregnancy, there was a measurable increase in luteal vascularization [184] and luteal blood flow increased from days 16 to 23 and again from days 24 to 50 [185]. Nitta et al. [178] demonstrated modulation of a lymph vessel formation marker and the VEGFs (VEGFC and VEGFD) responsible for lymph vessel formation in the CL of pregnancy. In vitro, IFNT upregulated VEGFC and ISG15 in both luteal endothelial cells and lymphatic endothelial cells and induced proliferation of lymphatic endothelial cells [178]. These data indicate that there may be modulation of both blood vessels and lymph vessels during early pregnancy, to support luteal survival. Possible changes in lymphangiogenesis are interesting in light of the altered immune cell populations in the CL during early pregnancy, discussed below. Interferons are antiviral cytokines that act on immune cells and alter their function. While it is possible that IFNT could act on the resident immune cells of the CL to alter their function during MRP, there is a dearth of information regarding specific actions of any conceptus-derived hormone on immune cells of the CL. Yankey et al. [186] were the first to report potential IFNT-regulated effects on peripheral immune cells when they reported upregulation of MX protein, an ISG, in PBMC of pregnant ewes compared with cyclic ewes. Since that first report, numerous others have reported upregulation of ISGs in peripheral immune cells in ewes [172,187] and cows [188,189]. Despite the extensive evidence that IFNT affects immune cells, little is known about the role that this upregulation of ISGs may play, either in the periphery or on the resident immune cells of the CL or uterus. In one of the only studies of IFNT effects on luteal cell-immune cell interactions, Shirasuna et al. [118] demonstrated that polymorphonuclear cells preincubated with physiological concentrations of IFNT were able to increase progesterone production in cultured luteal cells, while those that were not preincubated with IFNT did not. This may seem counterintuitive when considered in light of the earlier finding that neutrophil infiltration occurs early in luteolysis, as it is unlikely that neutrophils could have a luteotrophic and a luteolytic role [115]. Shirasuna and Miyamoto [190] suggest that neutrophils can be programmed into N1 (antitumoral) and N2 (protumoral) phenotypes that would compromise or support luteal function, respectively. Neutrophils are a short-lived cell type, so it is unlikely that they are programmed by the luteal microenvironment, but they may be programmed by interferon tau in the periphery prior to luteal infiltration.

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In multiple reports, either pregnancy [191] or IFNT treatment [192,193] induced expression of the regulatory cytokine, IL10, in bovine and murine PBMC or a macrophage cell line. IFNT induced another resolving-type cytokine, IL4, in murine lymphocytes [192] and reduced total proliferation and the number of proliferating CD8+ T cells in bovine mixed lymphocyte reactions [194]. In addition, in murine cells or a human cell line, IFNT reduced IL17 [195], IL6, TNF [193], IL1B, and phagocytic activity [196], all indicative of immune activation and inflammation. Overall, these findings indicate that IFNT may have some ability to generate a tolerogenic immune environment, which may have an effect on the uterus and ovary during MRP.

Luteal Rescue in Other Species Primate The mechanism by which the primate CL is rescued is quite different than the mechanism by which the CL of any domestic animal is rescued. A primate ovary will ovulate, form a CL, and undergo luteolysis normally in the absence of the uterus, meaning that there is no requirement for uterine prostaglandins to induce luteal regression in the primate. Therefore, typically the mechanism for luteal rescue in the primate has been referred to as a luteotropic mechanism, that is, the CL is directly supported by the conceptus. The chorion of the primate produces a gonadotropin, known in the human as hCG. This gonadotropin binds directly to the LHCGR and provides luteotropic support to the CL of pregnancy. Luteal rescue occurs in humans around day 10, after which luteal progesterone, estrogen, inhibin A, and relaxin in plasma increase in concentration, which is attributable to the increased function of the CL [197] The primate CL regresses during a normal cycle in the presence of physiological concentrations of pulsatile LH and requires hCG for luteal rescue. This difference in requirement may be due to the increased endogenous concentration of hCG compared with LH, as the CL of the later cycle becomes less and less LH responsive, or to differences in signaling by these two gonadotropins, or may encompass components of both. Luteal steroidogenic cells derived from a mid-luteal phase macaque CL were even more gonadotropin responsive than cells from a late-luteal phase CL [198], indicating lesser sensitivity to gonadotropin stimulation as the luteal phase progresses. CG is produced at rapidly increasing concentrations [199], perhaps to compensate for the reduced sensitivity of the later stage CL to gonadotropin stimulation. Exposure to LH or to hCG can have somewhat different functional outcomes for the CL. Either continuous infusion of hCG (in 2/3 monkeys), or exponentially increasing concentrations of either LH or hCG (in all

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monkeys), which more closely mimics the increasing concentrations of hCG seen in pregnancy, maintained the CL, but continuous infusion of LH failed to rescue the CL [200]. This indicates a functional difference in LH and hCG responses, despite the finding that these molecules bind to the same receptor [201]. Interestingly, LH and hCG can result in somewhat different cellular signaling events [202]. While both LH and hCG activate cAMP, in this study hCG activated cAMP more potently than did LH. Moreover, although both hormones also activated MAPK and AKT signaling pathways, LH was a more potent activator of these pathways than hCG [202]. This may be in part because of the more sustained signal that is derived from hCG as compared with LH [203]. As expected, global gene expression profiling demonstrated that steroid hormone biosynthesis-related genes increased in response to hCG [204]. This is supported by a study of structural changes in the rhesus monkey CL of early pregnancy, compared with the menstrual cycle, that found fewer lipid droplets in the cytoplasm of steroidogenic cells in pregnancy, likely resulting from the increase in steroidogenesis [205]. The primate CL of pregnancy does not experience any dramatic increase in vasculature [206], despite an increase in steroid and peptide hormone output that is likely greater than that in the ruminant. Rothchild [1] was the first to propose autoregulation of the CL by progesterone. In fact, it appears that autoregulation is a key component of luteal rescue in the primate. In the presence of trilostane, an inhibitor of HSD3B1, hCG was not able to rescue the CL and luteal weight declined [207], indicating that progesterone itself is an important component of luteal maintenance. Although the majority of interest in autoregulation of the CL by steroids has been in regard to progesterone [208], investigators have also suggested that estrogens and glucocorticoids may be involved in mediating autoregulation of the CL [209,210]. Inhibition of steroid production by trilostane altered gene expression, which was only partially restored by the synthetic progestin, R5020 [211]. Those transcripts that were regulated by trilostane but not by progestin replacement are potential candidates for regulation by other steroid hormones downstream of HSD3B1. Progesterone is an immune inhibitory steroid hormone [102,103], so one might hypothesize that one important component of progesterone autoregulation of the CL is generation of a microenvironment conducive to programming of luteal resident immune cells toward antiinflammatory or regulatory phenotypes. In most domestic animals, luteal progesterone is required for most, or all, of the length of pregnancy. However, the human and nonhuman primate only require luteal progesterone production for approximately the first 4 weeks of pregnancy. This, however, does not make the role of the primate CL any less instrumental in

pregnancy establishment. As in domestic animal pregnancies, failure of luteal rescue in the primate results in demise of the embryo and loss of the pregnancy. Rodents Luteal function of the rodent is quite different than that of domestic livestock or primates, in that there is not sustained luteal function during the diestrus phase of the natural cycle. Rather, during a cycle in which the rodent is not bred, the CL develops and secretes progesterone, but regresses within a day of formation. However, in the case that a mating occurs, the cervical stimulation from mating results in a diurnal and a nocturnal PRL surge that continue twice daily for 10–12 days following mating, even in the absence of a conceptus [212]. This PRL provides luteotropic support and allows the rodent CL to form a fully functional, sustained CL. A study of PRLR null mice demonstrated that important roles of PRL signaling included upregulation of LHCGR and thus support of steroidogenesis, and downregulation of AKR1C1 [213]. The CL of pseudopregnancy would be unlikely to form in the wild, unless embryonic loss followed mating, but it has been studied extensively in laboratory settings to gain a better understanding of luteal function in the rodent. The CL of pregnancy must be rescued after the first 10 days of pregnancy, during which it is supported by PRL. Following the period of luteal support by pituitary PRL, there are three important luteotropins in the rodent CL during pregnancy. Initially, during the period when the CL is supported by pituitary PRL, additional PRL from the decidua, which begins to be produced around day 7 in the rat, also provides luteotropic support [214]. Pituitary PRL declines rapidly around days 10–12 in the rat, after which the CL is supported by CSH1 and CSH2, which are produced by the giant cells of the trophoblast. CSH1 and CSH2 both have similar actions to PRL and bind to the PRL receptor; in an in vitro study of murine luteal cells, CSH1 and CSH2 elicited a similar steroidogenic response as PRL. CSH1 was greater in the trophoblast immediately following loss of pituitary support, while CSH2 was greater in late pregnancy, indicating a switch from CSH1 to CSH2 support as pregnancy progresses [215]. A study that compared the proteome of the functional CL of pregnancy to the regressing CL of pregnancy and the CL of lactation in the rat demonstrated similar expression of proteins related to steroidogenesis in the functional CL of pregnancy and lactation, but greater expression of antioxidant-type proteins during pregnancy, perhaps to protect luteal tissue from damage induced by reactive oxygen species produced during steroidogenesis. Interestingly, this study also demonstrated high concentrations of AKR1C1 in the regressing CL of pregnancy and low concentrations of the same enzyme in the growing CL of lactation, even as they are present

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on the ovary at the exact same time [216]. For a more details regarding the rodent CL of pregnancy, see Refs. [215,217]. For a comparison of mechanisms observed during luteal rescue in the ruminant, primate, and rodent, see Fig. 3. Other Species In the pig, the conceptus produces estrogens, particularly estradiol 17-beta, which acts to shift endometrial prostaglandin release from endocrine secretion during

the estrous cycle to exocrine secretion during pregnancy, thus sequestering PGF2A in the uterine lumen, rather than allowing it to escape the uterus and initiate luteolysis [218]. Very little is known about MRP in the horse [219]. The conceptus produces prostaglandins, which induce its movement throughout the uterus early in pregnancy. The migration of the embryo throughout the uterus is necessary for luteal rescue. In the dog, unlike in humans or any other domestic species, the CL of the estrous cycle and the CL of pregnancy are equally long lived, so luteal rescue is unnecessary [220]. FIG. 3 Mechanisms for rescue of the corpus luteum in early pregnancy in ruminants, primates and rodents. (A) In the ruminant, IFNT is produced by the conceptus and exerts effects on the CL, the uterus, and peripheral blood cells. In the CL, IFNT increases luteal resistance to the luteolytic effects of PGF2A, perhaps in part by reducing expression of THBS1, SERPINE1, TGFB1, and EDN1 and by increasing cell survival. IFNT also induces VEGFC, a marker of lymphangiogenesis. IFNT exerts an antiluteolytic effect on the uterus, reducing expression of OXTR via ESR1 and therefore reducing OXT-induced PGF2A pulses, but not basal PGF2A release from the uterus. IFNT also increases the ratio of PGE2:PGF2A and may reduce prostaglandin transport to the CL, by decreasing expression of the prostaglandin transporter protein, PGT. In peripheral immune cells, IFNT induces ISGs and IL10 expression and reduces lymphocyte proliferation. Neutrophils treated with IFNT gain the ability to induce progesterone production by cultured luteal cells, perhaps because they become an N2, protumoral, phenotype. (B) In the primate, as the CL becomes more resistant to the actions of LH (dashed line), the chorion of the placenta produces high concentrations of hCG, which bind to the LH receptor (LHCGR), increase intracellular cAMP, and lead to an increase in steroid biosynthesis genes and steroidogenesis. (C) In the rodent, PRL is released from the anterior pituitary in response to mating stimulus in diurnal and nocturnal pulses and is necessary to support luteal function early in pregnancy. The decidua produces PRL around day 7, further supporting luteal maintenance. Late in pregnancy, CSH1 and CSH2, produced by trophoblast giant cells, become the most important luteotropic signal. Signaling downstream of the PRLR includes upregulation of the LHCGR, increased steroidogenesis, downregulation of AKR1C1 to decrease progesterone catabolism, and ultimately, luteal survival.

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Other Changes in the CL During Early Pregnancy Since Lobel and Levy [87] first reported the presence of immune cells in the CL and lesser infiltration of these cells into the CL of early pregnancy, evidence has accrued for immunological modulation in the CL during early pregnancy. Whether these changes are due to IFNT exposure, to another conceptus secretion, or to another factor remains to be determined. Luteal resident immune cell phenotype is altered in the CL of pregnancy, compared with the CL of the estrous cycle, which indicates a likely change in function. Poole and Pate [92] reported that luteal T cell phenotype is altered in early pregnancy, with a greater number of CD8αβ+ T cells in the CL of pregnancy than in the CL of the estrous cycle. Interestingly, expression of the CD8αβ heterodimer is induced particularly on gamma delta T cells in the CL of early pregnancy, indicating changes in the function of these cells. Therefore, immune cells are not simply repressed in the CL of pregnancy, but are modulated, perhaps to protect the CL from regression. Interestingly, MHCIA, the antigen presentation molecule that presents antigen to cytotoxic T cells and is expressed on most or all cells of the body, was 15-fold greater in the CL of days 20–25 of pregnancy than in the CL of the estrous cycle [221], although this does not provide any indication of whether antigen presentation was different comparing the estrous cycle to pregnancy. Global gene profiling studies also support the idea that immune function is modulated in the CL during early pregnancy and may be imperative for luteal rescue. In a study of the ovine CL during the late cycle, early regression, and early pregnancy, modulation of immunerelated genes was evident, with differentially abundant mRNA, including PTX3 and IL6. A predicted modulated function, comparing CL of the cycle and CL of pregnancy was related to cell adhesion and chemokines [173]. Cytokine and chemokine transcript abundance also change in the CL as pregnancy progresses. Eotaxin, which recruits eosinophils, had increased transcript abundance as pregnancy progressed, with a peak on days 150–160 of pregnancy. Conversely, lymphotactin, which recruits T cells, decreased during this same time period. These data indicate that immune cell populations may be dynamic in the CL throughout pregnancy [221]. The leporine CL of pseudopregnancy and the regressing CL at parturition both have a greater number of macrophages than observed in the CL of pregnancy. Regressing CL and CL of pseudopregnancy produced TNF in response to lipopolysaccharide stimulation, while the CL of pregnancy did not [121]. MHCII expression, a marker of ability to present antigen to immune cells, was lesser during early pregnancy than on the same day of the cycle in cows (day 18) [222], indicating a cellular environment with altered communication to immune cells in early pregnancy.

As in the ruminant CL, the primate CL of early pregnancy seems to undergo modulation of immune function. Similarly, LH ablation/replacement studies and early pregnancy simulation by hCG treatment indicated that there are immune-type functions that are regulated by LH and hCG in the primate CL [204,211]. These gonadotropins are not global immune inhibitors, because the pathway analysis indicated that hCG upregulated some pathways but downregulated others [204]. Cocultures of human luteal cells with allogeneic peripheral blood mononuclear cells produced more progesterone and more IL4 and IL10 than either cell type alone, with the greatest effect observed when the luteal cells were from the CL of pregnancy [95] indicating a trophic relationship between these two cell types. Together, these data indicate that immune regulation in the CL during MRP, and as pregnancy progresses, is complex. Rather than immune activation being either turned on in the case of regression or turned off in the case of luteal rescue, resident immune cells are differentially regulated to achieve these outcomes. Intraluteal prostaglandins may also play a role in protecting the CL from luteolysis. In a study that compared prostaglandin synthesis in the CL of pregnant or nonpregnant ewes after treatment with PGF2A, the ratio of AKR1C3 (which is responsible for synthesis of PGF2A) to PTGES (which is responsible for synthesis of PGE2), found that PGE2 synthesis was likely to be greater in the CL of pregnancy. Furthermore, the in vitro metabolism of PGF2A to PGFM was greater in the CL of day 14 of pregnancy than in the CL of day 14 of the cycle [223]. Indeed, prostaglandin production, metabolism, and signaling are altered in the CL of pregnancy. In ovarian venous plasma, greater concentrations of PGF2A were present on D14 of the estrous cycle, but greater concentrations of PGFM and PGE2 were present on D14 of pregnancy [167]. Given that PGE2 is generally thought of as a luteotropic prostaglandin that supports luteal survival and progesterone production, these data may indicate that the balance of intraluteal prostaglandin production and metabolism is a key player in luteal rescue. A recent miRNA profiling study demonstrated modulation of 12 known and 3 novel miRNA in the CL during MRP. miRNA are small (20–22 nucleotides) inhibitors of translation. They specifically degrade or inhibit the translation of their targets, based on sequence complementarity. Analysis of the predicted targets of these miRNA indicated that they are involved in functions such as immune regulation and regulation of apoptosis [224]. Prosurvival pathways are upregulated in the CL of pregnancy, while death pathways are upregulated in the CL of the cycle [225]. The data from Maalouf et al. [224] indicated that this regulation of survival and death may be regulated by miRNA in the CL.

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CONCLUSION

Additional Potential Regulators of the CL in Early Pregnancy The conceptus has many other secretory products in addition to IFNT. In an in vitro experiment, IFNT did not protect cultured ovine LLC from a PGF2Ainduced reduction in progesterone, but IFNT-depleted conceptus secretory products, derived from days 15 to 16 conceptuses, did [226], indicating that, although IFNT is the only antiluteolytic conceptus secretory product in the ruminant [155,157] it is not the only conceptus secretory product that may affect luteal function. Further, despite the finding that in vitro IFNT treatment does not induce progesterone production compared with control [118], the bovine CL of day 28 of pregnancy secretes more progesterone than the CL of day 14 of the cycle in vitro [227] and luteal tissue slices from ovine CL of a normal pregnancy secreted more progesterone than CL of the cycle or CL derived from ewes carrying an abnormal or unhealthy embryo [228], indicating that a healthy pregnancy may support increased luteal progesterone production. PAG are produced in abundance by the binucleate giant cells of the placenta, beginning during the third week of pregnancy [229], making them unlikely to have been the luteoprotective proteins observed by Wiltbank et al. [226]. However, PAG are present in the circulation at high concentrations and may have an important signaling role following the MRP period. Ovine luteal tissue slices from pregnant animals produce greater concentrations of progesterone, more PGE2 and arachidonic acid in response to PAG, than those from cyclic animals, but failed to respond to LH as did those from cyclic animals [230]. Similar responses have been demonstrated in the cow [231–233], indicating that in the ruminant, there may be a switch from hypophyseal LH to luteal PGE2 as the primary luteotropin in later pregnancy, with PAG as important supporters of local PGE2 production.

Another Pivotal Maternal Recognition Period? In ruminants, an additional phenomenon worth noting is that of continued luteal maintenance. Although IFNT may alter PGF2A release from the uterus and/or render the CL less sensitive to the actions of PGF2A, IFNT secretion peaks at day 23 of pregnancy in the cow and declines thereafter [234]. This raises the question: what is responsible for continued luteal maintenance? A recent large-scale study on a commercial dairy farm was used to investigate maintenance of the CL that was contralateral to the conceptus-bearing uterine horn. The contralateral CL was induced with GnRH on day 5 after estrus and artificial insemination and regressed in two clearly defined groups. Approximately 25% of contralateral CL regressed on days 19–25, as expected (likely as a

result of failure of IFNT to reach the horn contralateral to the pregnancy and exert its antiluteolytic effect), while 75% of contralateral luteal regressions occurred between days 33 and 60 of pregnancy, later than expected. These data indicate that there is a second period of luteal rescue in cattle, around days 33–60, and that this second period also relies on a local mechanism for either alteration of PGF2A release or for delivery of a luteoprotective factor, because in most cows that underwent contralateral luteal regression, the ipsilateral CL was maintained [235]. Interestingly, the timing of this second period of luteal regression corresponds to the timing of observation of the embryonic vesicle growing large enough to reach the contralateral horn of the uterus (days 31–45) [89]. Perhaps, there is an important local signaling event that takes place between the growing embryo and the uterus or CL, after the classical MRP period, to prevent regression. Similarly, embryonic loss prior to the onset of PAG production did not extend luteal lifespan, but when embryonic loss occurred after the onset of PAG production, there was an extended period of luteal maintenance [236]. This indicates that IFNT signaling alone was inadequate to rescue the CL, when the embryo died prior to the onset of PAG production. Perhaps, the exposure to IFNT was not prolonged enough or IFNT concentration was not great enough to rescue the CL or perhaps luteal rescue has multiple components, not limited to IFNT, and PAG exposure to the uterus or CL is required for continued suppression of luteolysis.

CONCLUSION In the nearly three and a half centuries since the CL was first observed and described in detail, as a transient yellow globule on the ovary, this endocrine gland has intrigued scientists and physicians alike. The transience of the CL is intimately linked to its physiological functions, although the mechanisms regulating both luteolysis and luteal survival are diverse. Understanding the functions and endocrine and paracrine secretions of the various cell types of the CL, including immune and endothelial cells, have led to a complex understanding of the cell-cell communication that is required for luteal regression to occur. Although our basic understanding of luteal function is built on important early findings in the 1970s and 1980s, new technologies, including big data approaches such as transcriptomics and proteomics, have improved our understanding of luteal regression and luteal rescue. Much is known about luteal function, but gaps remain in our understanding. Given that proper luteal function is obligatory for fertility and reproduction, luteal research will remain a priority for both clinical and basic researchers.

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[225] Lee J, Banu SK, McCracken JA, Arosh JA. Early pregnancy modulates survival and apoptosis pathways in the corpus luteum in sheep. Reproduction 2016;151:187–202. [226] Wiltbank MC, Wiepz GJ, Knickerbocker JJ, Belfiore CJ, Niswender GD. Proteins secreted from the early ovine conceptus block the action of prostaglandin F2A on large luteal cells. Biol Reprod 1992;46:475–82. [227] Wickersham EW, Tenabe TY. Functional state of bovine corpora lutea as determined by denovo production of progesteron in vitro, J Anim Sci 1967;26(1):158–62. Available from: http:// www.ncbi.nlm.nih.gov/pubmed/8562708. [228] Abecia JA, Forcada F, Zúñiga O. Differences in reproductive performance, embryo development, interferon-tau secretion by the conceptus and luteal function in ewe lambs synchronized in oestrus before or after the spontaneous onset of luteal activity preceding puberty. Reprod Domest Anim 2001;36(2):73–7. [229] Wallace RM, Pohler KG, Smith MF, Green JA. Placental PAGs: gene origins, expression patterns, and use as markers of pregnancy. Reproduction 2015;149:R115–26. [230] Weems YS, Kim L, Humphreys V, Tsuda V, Blankfein R, Wong A, et al. Effect of luteinizing hormone (LH), pregnancy-specific protein B (PSPB), or arachidonic acid (AA) on ovine endometrium of the estrous cycle or placental secretion of prostaglandins E2 (PGE2) and F2alpha (PGF2alpha) and progesterone in vitro. Prostaglandins Other Lipid Mediat 2003;71:55–73. [231] Del Vecchio RP, Sutherland WD, Sasser RG. Effect of pregnancyspecific protein B on luteal cell progesterone, prostaglandin, and oxytocin production during two stages of the bovine estrous cycle. J Anim Sci 1995;73:2662–8. [232] Del Vecchio RP, Sutherland WD, Sasser RG. Bovine luteal cell production in vitro of prostaglandin E2, oxytocin and progesterone in response to pregnancy-specific protein B and prostaglandin F2A. J Reprod Fertil 1996;107:131–6. [233] Weems YS, Lammoglia MA, Vera-Avila HR, Randel RD, Sasser RG, Weems CW. Effects of luteinizing hormone (LH), PGE2, 8-EPI-PGE1, 8-EPI-PGF2A, trichosanthin and pregnancy specific protein B (PSPB) on secretion of prostaglandin (PG) E (PGE) or F2A (PGF2A) in vitro by corpora lutea (CL) from nonpregnant and pregnant cows. Prostaglandins Other Lipid Mediat 1998;55:27–42. [234] Stojkovic M, Wolf E, Btittner M, Berg U, Charpigny G, Schmitt A, et al. Secretion of biologically active interferon tau by in vitroderived bovine trophoblastic tissue. Biol Reprod 1995;53: 1500–7. [235] Baez GM, Tresvisol E, Barletta RV, Cardoso BO, Ricci A, Guenther JN, et al. Proposal of a new model for CL regression or maintenance during pregnancy on the basis of timing of regression of contralateral, accessory CL in pregnant cows. Theriogenology 2017;89:214–25. [236] Wijma R, Stangaferro ML, Kamat MM, Vasudevan S, Ott TL, Giordano JO. Embryo mortality around the period of maintenance of the corpus luteum causes alterations to the ovarian function of lactating dairy cows. Biol Reprod 2016;95(112):1–14.

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18 Transgenic Mouse Models in the Study of Ovarian Function Amanda Rodriguez, Robert T. Rydzea, Shawn M. Brileya, Stephanie A. Pangasa Abbreviations AKT Aldh1a1 Aldh1a2 ALK2 ACVR2 Amh Amhr2 Aqp8 Areg Atg7 Bax Bcl2 Bclx Becn1 Bmp cAMP Casp Cdc25 Cdk1 cGMP CL COC Cox2 Cx37 Cxcr4 DAZL DMRT1 E# EGF Emx2 Erα Erβ Ereg Fanca Fance Figla Fog2 Foxc1

a

protein kinase B aldehyde dehydrogenase 1 family member A1 aldehyde dehydrogenase 1 family member A2 activin A receptor type 1 activin A receptor type 2A anti-M€ ullerian hormone anti-M€ ullerian hormone type 2 aquaporin 8 amphiregulin autophagy-related 7 CL2 associated X, apoptosis regulator Bcl2, apoptosis regulator Bcl2 like 1 Beclin-1 bone morphogenetic protein cyclin adenosine monophosphate caspase cell division cycle 25 cyclin-dependent kinase 1 cyclin guanosine monophosphate corpora lutea cumulus-oocyte complex prostaglandin-endoperoxide synthase 2 connexin 37 C-X-C motif chemokine receptor 4 deleted in azoospermia like doublesex and Mab3-related transcription factor 1 embryonic day epidermal growth factor empty spiracles homeobox 2 estrogen receptor alpha estrogen receptor beta epiregulin Fanconi anemia complementation group A Fanconi anemia complementation group E folliculogenesis-specific BHLH transcription factor zinc finger protein, FOG family member 2 forkhead box C1

Foxl2 Foxo3a FSH Fst Gas2 Gata4 GCC Gdf9 Gja1 Grem1 GV GVBD γH2AX hCG Hes1 IGF1 Inha Inhb Inhbb Kit KL KO Lats1 LH Lhx1 Lhx8 Lhx9 Lkb1 Mapk Mpf mTORC Nanog Nobox Nos1 Nos2 Nos3 Nppc Npr2 Pde3a Pdk1

forkhead boxl2 forkhead box 03 follicle stimulating hormone follistatin growth arrest specific 2 GATA-binding protein 4 germ cell cyst growth differentiation factor 9 gap junction protein alpha1 gremlin 1 germinal vesicle germinal vesicle breakdown H2A histone family member X human chorionic gonadotropin Hes family BHLH transcription factor 1 insulin-like growth factor 1 inhibin beta A subunit inhibin B inhibin beta B subunit KIT proto oncogene receptor tyrosine kinase kit ligand knockout large tumor suppressor kinase 1 luteinizing hormone lim homeobox 1 lim homeobox 8 Lim homeobox 9 liver kinase b1 mitogen activated protein kinase maturation promoting factor mechanistic target of rapamycin nanog homeobox newborn ovary homeobox encoding nitric oxide synthase 1 nitric oxide synthase 2 nitric oxide synthase 3 natriuretic peptide C natriuretic peptide receptor 2 phosphodiesterase 3a pyruvate dehydrogenase kinase 1

These authors contributed equally.

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296 PGC Pgs PI3K PKA PMSG PN# POI Pou5f1 PR Prdm1 Prdm14 PTEN Ptger2 Robo1 Rps6 Rptor Rspo1 S6k Sdf1 Sf1 Sohlh1 Sohlh2 Stra8 Taf4b TGFβ TNAP TNFIP6 TSC1 TSC2 Wee1b Wnt4 WT1 Zfx Zp1 Zp2 Zp3

18. TRANSGENIC MOUSE MODELS IN THE STUDY OF OVARIAN FUNCTION

primordial germ cell prostaglandin endoperoxide synthase 2 phosphatidylinositol-4,5-bisphosphate 3-kinase protein kinase A pregnant mare’s serum gonadotropin postnatal day # primary ovarian insufficiency POU class 5 homeobox 1 progesterone receptor PR/Set domain 1 PR/Set domain 14 phosphatase and tensin homolog prostaglandin E receptor 2 roundabout guidance receptor 1 ribosomal protein s6 regulatory associated protein of mTOR complex 1 R-spondin1 ribosomal protein s6 kinase stromal cell derived factor 1 splicing factor 1 spermatogenesis and oogenesis specific basic helix loop helix 1 spermatogenesis and oogenesis specific basic helix loop helix 2 stimulated by retinoic acid 8 TATA-box-binding protein associated factor 4b transforming growth factor beta tissue nonspecific ALP TNF alpha induced protein 6 tuberous sclerosis 1 tuberous sclerosis 2 wee1 G2 checkpoint kinase wingless-type MMTV integration site family, member 4A Wilms tumor 1 zinc finger protein, X-linked zona pellucida 1 zona pellucida 2 zona pellucida 3

INTRODUCTION The mammalian ovary is a highly organized, complex organ containing the germ cells (oocytes) and major somatic support cells: granulosa, thecal, and stromal cells. Germ cells are maintained within a quiescent pool of small ovarian follicles called primordial follicles (Fig. 1). The primordial follicles make up what is termed the true “ovarian reserve,” the size of which is thought to ultimately dictate the female reproductive lifespan. Disruptions in key processes controlling ovarian differentiation, oocyte maturation, folliculogenesis, and hormone production ultimately impair an oocyte’s ability to be fertilized and cause infertility. Advancements in transgenic mouse models have been vital for identifying critical genes and the pathways in which they function during ovarian development. Over the past 30 years, the use of tissue and cell-specific genetic recombination, such as the cre recombinase-loxP system, has served as a sophisticated genetic tool for generating mutant mouse models (Table 1), as has more traditional knockout (KO) and knockin mouse genetic models. More modern

techniques, such as those using CRISPR-Cas9 will further revolutionize our ability to manipulate the genome and test the roles of specific mutations. In this chapter, we review how genetically engineered mice have served to reveal the mechanisms behind various forms of ovarian dysfunction and female infertility. Often, these mechanisms are controlled by conserved developmental protein families, such as the transforming growth factor beta (TGFβ), wingless-type MMTV integration site (WNT), phosphatidylinositol-4,5-bisphosphate 3-kinase (PI3K)/ phosphatase and tensin homolog (PTEN)/thymoma viral proto-oncogene (AKT), and Notch, as they function in key events: primordial germ cell (PGC) specification, follicular growth, meiosis, and ovulation.

OVARIAN DIFFERENTIATION AND GERM CELL DEVELOPMENT In mammals, PGCs form early during development outside of the embryo proper at the time of gastrulation and must find their way to the gonadal primordium (gonadal or genital ridge) located on the ventral surface of the mesonephros. From just a few cells ( 3.4 ng/mL During COS

- Development of 19 follicles - Serum E2 > 3500 pg/mL After ovum pick-up

- Retrieval >15 oocytes has been considered a good predictor of OHSS, with a sensitivity of 90.5% and specificity of 81.3% [69]. Levels >10 ng/mL are associated with a threefold increase in the incidence of OHSS [70]. Antral follicle count (AFC) is also predictive of OHSS [68,71]. The risk of OHSS increases from 2.2% to 8.6% in women with an AFC 24 [61].

During COS RISK FACTORS In theory, OHSS could develop in any patient undergoing COS with gonadotropins. However, the reality is that there are some women who are at a much higher risk. Identifying these women is essential for lowering and even eliminating OHSS in clinical practice [59] (Table 1).

Before COS Begins Not much can be said about the phenotype. However, ovulatory disorders, particularly PCOS, are the most prevalent characteristic of women at risk of OHSS. Similarly, women who have previously developed OHSS should be considered at high risk of experiencing a similar clinical picture [60–65]. Black women also appear to be at higher risk of OHSS [60]. In addition, younger age (20 follicles during COS significantly increases the risk of OHSS [75]. In fact, a model has been developed to predict OHSS with 82% sensitivity and 90% specificity: >19 large-/medium-sized follicles before ovulation triggering; >25 follicles at oocyte retrieval; and >24 oocytes retrieved [73]. Serum estradiol concentrations >3500 pg/mL are also associated with OHSS [62–64,66–69,78,79].

After Ovum Pickup The number of oocytes retrieved is the most direct measure of the ovarian response, although in some cases there are difficulties in obtaining eggs. Retrieval of >15 oocytes significantly increases the chance of OHSS in cycles in which hCG has been used to trigger ovulation [72].

CLINICAL MANIFESTATIONS The variety of clinical manifestations of OHSS is the consequence of the processes that define the syndrome

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CLINICAL MANIFESTATIONS

(Fig. 1). Enlarged ovaries may themselves produce abdominal discomfort. Increased ovarian vascular permeability leads first to fluid accumulation in the abdomen and other body cavities, which then leads to abdominal heaviness and breathing difficulties due to limited diaphragmatic mobility [80]. Furthermore, this shift in serum from the intravascular to the extravascular space causes haemoconcentration and reduced blood perfusion, resulting in reduced general organ perfusion. Oliguria and renal insufficiency may occur, and liver function may also be affected. Moreover, haemoconcentration increases the risk of thromboembolic events. In very severe forms, renal failure and reduced perfusion in other vital organs, such as the brain and heart, may have fatal consequences [81]. There are two clinical forms of OHSS, both hCG related: the early onset, occurring 3–7 days after hCG administration; and the late onset, occurring 12–17 days after hCG administration, which is related to pregnancy-induced hCG production [82]. The early onset is usually mild to moderate, while late OHSS is more severe as the rising hCG during pregnancy exacerbates the course of the syndrome. In order to define an increasing degree of severity in the establishment of OHSS, different classifications have been published [83,84] based upon the severity of symptoms, signs, and laboratory findings. OHSS classification is employed for academic reasons, but is also used as a guideline to establish who should receive outpatient management and who should be hospitalized. A widely employed classification is described in Table 2 [85]. Here, mild OHSS is characterized by bilateral ovarian enlargement with multiple follicular and corpus luteum cysts, abdominal distention and discomfort, mild nausea, and less frequently, vomiting and diarrhea. There are no biochemical abnormalities. No special care is necessary, but surveillance of the patient is indicated. The clinical features of moderate OHSS include ultrasonographic evidence of ascites. Ovaries are frequently enlarged up to 12 cm. Abdominal discomfort and gastrointestinal symptoms such as nausea, vomiting, and diarrhea are more frequent and intense than in mild OHSS. A sudden increase in weight of >3 kg (6.6 lb) might be an early sign of moderate OHSS (Table 2). Laboratory features include a hematocrit >41% and white blood cell concentration (WBC) >15,000/mL along with hypoproteinemia. To consider OHSS as severe, clinical evidence of ascites with severe abdominal pain and, in some patients, pleural effusion is pathognomonic. Ascites and pleural effusion may compromise pulmonary function, resulting in hypoxia (Table 2) [86]. Women with severe OHSS can gain as much as 15–20 kg (33–44 lb) over 5–10 days and display progressive leukocytosis. Hypovolemia, oliguria,

TABLE 2 OHSS Stage

Clinical Classification of OHSS (Adapted From Ref. [85].) Clinical Features

Laboratorial Features

Mild

Abdominal distention/ discomfort Mild nausea/vomiting Diarrhea Enlarged ovaries

No main laboratorial alterations

Moderate

Mild features + ultrasonographic evidence of ascitis

Elevated hematocrit (>41%) Elevated WBC (>15,000) Hypoproteinemia

Severe illness

Mild + moderate features +: Clinical evidence of ascitis Hydrothorax Severe dyspnea Oliguria/anuria Intractable nausea/vomiting Tense ascitis Low blood/central venous pressure Rapid weight gain (>1 kg in 24 h) Syncope Severe abdominal pain Venous thrombosis

Hemoconcentration (Htc > 55%) WBC > 25,000 Creatinine clearance 1.6 mg/dL Hyponatremia (Na+ < 135 mEq/L) Hypokalemia (K+ < 5 mEq/L) Elevated liver enzymes

Critical

Anuria/acute renal failure Thromboembolism Arrhythmia Pericardial effusion Massive hydrothorax Arterial thrombosis Adult respiratory distress syndrome Sepsis

Worsening of severe findings

or anuria, and intractable nausea and/or vomiting are frequently present. Creatinine levels are >1.6 mg/dL. Reduced liver perfusion results in the depletion of anticlotting factors and transaminases are increased [87]. Other laboratory findings include a hematocrit >55%, WBC count >25,000/ mL, hyponatremia, and hyperkalemia [2,88,89]. Hemoconcentration increases the risk for thromboembolism. In critical OHSS, the function of vital organs and systems is seriously compromised. A series of catastrophic events could occur presenting a life-threatening situation. First, OHSS can be complicated with hemoconcentration, with a risk of venous (75%) and arterial (25%) thrombosis that may lead to permanent neurological injury or death [90,91]. High levels of factor V, platelets, fibrinogen, profibrinolysin, fibrinolytic inhibitors, and increased thromboplastin generation are observed in women with OHSS [92]. Moreover, altered liver function can lead to disseminated intravascular coagulation (DIC) and liver failure with hepatic encephalopathy [87].

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Thromboembolic complications of OHSS have been reported in the internal jugular, subclavian, axillary, and mesenteric vessels [86,93]. Cerebrovascular thrombosis is typically present as an ischemic infarct. In 40% of cases, these complications are associated with an underlying thrombophilia [94]. It is important to rule out thrombophilia before COS begins in order to consider low-dose heparin prophylactic therapy [95]. Other catastrophic events characteristic of severe OHSS include acute kidney injury with anuria, sepsis and acute respiratory distress syndrome (ARDS), and cerebral edema/acute ischemia/encephalopathy [2,3,84,96–98]. Rarely, OHSS results in death. The precise risk of mortality from OHSS is unknown, but ranges between 0 and 3/100,000 in some reports [3,99,100]. Other potential complications of OHSS include ovarian torsion and hemorrhage from ovarian rupture. Ovarian torsion is the potential consequence of increased ovarian volume. It is characterized by ovarian enlargement, abdominal pain, nausea, vomiting, hypotension, progressive leukocytosis, and anemia. Ovarian torsion may demand surgical correction [101].

DIAGNOSIS The diagnosis of OHSS is based on clinical features (Table 3) [102]. The symptoms of OHSS are not specific and there are no diagnostic tests for the condition. Although patients may come to the clinic with abdominal distension and discomfort following ovulation triggering, the typical patient appears 2–3 days after oocyte retrieval with abdominal discomfort and gastrointestinal symptoms such as nausea, vomiting, and diarrhea and perhaps reduced urinary output and/or constipation. Therefore, it is important to get an idea of the severity of the suspected OHSS in the initial interview by asking whether the patient has undergone COS recently, if she has experienced OHSS before, if she has been diagnosed with PCOS, the number of eggs retrieved, if embryo replacement was performed, and the time of presentation following ovulation triggering. This is important because early onset is usually self-limited without many complications, while late onset could be associated with critical events. A series of exams should be performed at this point (Table 3) to determine the severity of the syndrome (Table 2) and whether the patient should initially be managed as an outpatient, remain hospitalized, or even be admitted to an intensive care unit (ICU). There is not obvious and rigorous compartmentalization of patients, and the decisions should be made considering signs, symptoms, laboratory and image testing, as well as the special characteristics of the patient, including her

TABLE 3

Relevant History, Symptoms, and Exams to be Taken in a Woman With Suspected OHSS Development (Adapted From Ref. [102], With the Permission of the Royal College of Obstetricians and Gynecologists.)

History Diagnosis of PCOS Previous episode of OHSS Time of onset of symptoms relative to trigger of ovulation Medication used for trigger ovulation (hCG or GnRH agonist) Number of follicles developed on final monitoring scan Number of oocytes retrieved Embryo replacement performed? Symptoms Abdominal bloating Abdominal discomfort/pain, need for analgesia Nausea and vomiting Breathlessness, inability to lie flat or talk in full sentences Reduced urine output Leg swelling Vulvar swelling Associated comorbidities such as thrombosis Examination General: assess for dehydration, edema (pedal, vulvar, and sacral); record heart rate, respiratory rate, assess for pleural effusion, blood pressure, body weight Abdominal: assess for ascites, palpable mass, peritonism; measure girth Laboratory and imaging tests Full blood count Hematocrit C-reactive protein Urea and electrolytes (Na, K) Serum osmolality Liver function tests Coagulation profile D-dimers hCG (if embryo replacement) Ultrasound scan: ovarian size, pelvic, and abdominal free fluid Chest X-ray Other tests that may be indicated (in critical OHSS) Arterial blood gases Electrocardiogram (ECG)/echocardiogram Computerized tomography pulmonary angiogram (CTPA) or ventilation/perfusion (V/Q) scan

sensitivity to pain, her compliance to treatment and difficulty accessing the clinic if her condition worsens. Since OHSS symptoms are not specific, care must be taken to exclude other serious conditions that may be present in a similar manner but require very different management. The most frequent is peritonism, which is associated with some hemoperitoneum induced by blood loss from punctured ovaries. While OHSS is associated with elevated hematocrit and reduced serum osmolality and sodium, hemoperitoneum is accompanied by low hematocrit. Pyrexia is not typically present in OHSS,

V. HUMAN OVARIAN PATHOPHYSIOLOGY: SELECT ASPECTS

PREVENTION

and therefore pelvic infections, including appendicitis or bowel perforation after blind transvaginal oocyte retrieval, must be ruled out in the case of elevated temperature. Ovarian torsion is associated with progressive leukocytosis and anemia, and should also be considered a potential differential diagnosis. If the pregnancy test is positive, a rare complication could be an ectopic pregnancy from the preceding cycle, which should also be kept in mind.

PREVENTION Before COS Begins Planning COS in patients at risk is the most important step in the prevention of OHSS [103]. The characteristics defined in Table 1, which identify women at high risk of developing OHSS, should be carefully considered in order to avoid further complications. Selecting the Stimulation Protocol There are different reasons to recommend stimulation protocols utilizing GnRH antagonists for ovulation suppression, namely similar pregnancy rates to protocols employing long-GnRH agonist suppression, lower costs, and lower risk of developing OHSS. This appears to act at two levels: lower serum levels of estradiol during ovarian stimulation, and the possibility of triggering ovulation with a GnRH agonist by taking advantage of the flareup mechanism [104,105]. In fact, a main issue for the generalized use of pituitary suppression with an antagonist was lower pregnancy rates, which is probably associated with the lack of experience with GnRH antagonists. Today, there are reassuring data from 29 randomized clinical trials (RCTs) showing that GnRH antagonist protocols display similar pregnancy rates as long-agonist protocols. Moreover, they employ a smaller amount of gonadotropins, which reduces the costs of the cycle, and are also associated with less incidence of OHSS [106]. The use of a GnRH antagonist protocol, per se, reduces the risk of OHSS compared with protocols that use a GnRH agonist, most likely due to a reduction in circulating estradiol levels seen with GnRH antagonist suppression. Large randomized studies using long-agonists versus antagonists protocols in ART triggering ovulation with hCG have shown a significantly reduced incidence of severe OHSS in the antagonist group (5.1%) compared with the agonist (8.9%) [107]. Furthermore, in women with PCOS, it was observed that suppression with antagonist as opposed to agonist also reduced the incidence of moderate OHSS from 60% to 40% after ovulation triggering with hCG [108].

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It is also worth to stress that the dose of gonadotropins to be employed in women at risk should not be higher than 150 IU/day. Another possible approach is to substitute GnRH antagonists with progestogens in COS with cycle segmentation. No RCT has been established to test the role of these stimulation protocols in preventing OHSS, but they are widely employed and the published experience using 4–10 mg/day medroxyprogesterone acetate simultaneously with gonadotropins shows good-quality oocytes and pregnancy rates without LH surges or OHSS [109,110]. Finally, in vitro maturation (IVM) can also be considered in these patients. When IVM was described, avoidance of OHSS was always mentioned as one of the main advantages [111]. Monitoring PCOS patients after spontaneous or induced menses and administering 10,000 IU hCG when endometrial thickness is >6 mm has been associated with high yields of oocytes (>15), as well as acceptable implantation and pregnancy rates (30% and 50%, respectively) without OHSS development [112]. IVM remains unpopular, however, because results have not been replicated in many centers. Adjuvant Therapies ASPIRIN

Increased VEGF levels in women at risk of developing OHSS results in platelet activation and release of histamine, serotonin, platelet-derived growth factor, and/or lysophosphatidic acid, which can further potentiate the physiologic cascade of OHSS. Based on this theory, aspirin has been considered a possible therapy in OHSS [113]. In one set of randomized studies, the incidence of severe OHSS was reduced (1.7% vs 6.5%) in patients who received a daily dose of 100 mg aspirin plus prednisolone in varying doses (10–30 mg) from the first day of stimulation until the day of the pregnancy test [114]. In another study including women at high risk of OHSS, 100 mg aspirin/day were administered from the first day of the cycle (during COS) until the next menses in the case of a negative ART outcome or the ultrasonographic detection of embryonic cardiac activity. Aspirin significantly reduced the incidence of OHSS from 8.4% to 0.25% compared with no treatment [113]. METFORMIN

Metformin may improve intraovarian hyperandrogenism, and it can affect the ovarian response by reducing the number of nonperiovulatory follicles, thereby reducing estradiol secretion. By employing 500 mg three times daily or 850 mg twice daily during COS in PCOS patients, the incidence of OHSS was significantly reduced from 20.4% to 3.8% [115]. A meta-analysis including 12 studies showed that OHSS risk was significantly lower

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with metformin use (relative risk (RR) 0.44, 95% CI: 0.26–0.77), while maintaining the same ART outcomes as controls without metformin [116]. Apparently, metformin works better in obese than in nonobese patients [117].

During COS Coasting Coasting consists of withholding the administration of gonadotropins at the end of COS for up to 4 days to decrease the risk of OHSS. Although extensively employed in the past, several studies have shown that coasting does not decrease the risk of OHSS, but is associated with fewer oocytes retrieved [118], and prolonged coasting might even hurt oocytes quality and reduce implantation [119]. Ovulation Triggering With a GnRH Agonist hCG has been widely employed to mimic the midcycle surge of LH to trigger final oocyte maturation and ovulation for more than 60 years. hCG has a longer half-life than LH and it is known to initiate the cascade of events leading to OHSS. hCG stimulates VEGF release in granulosa-lutein cells that bind to VGEF-R2, which increases vascular permeability in the ovaries [11,32]. Administration of a GnRH agonist is associated with an initial “flare-up” effect in which both serum LH and FSH are increased [104]. It has been shown that a so-called GnRH agonist-induced surge of gonadotropins can last for 24–36 h and induce oocyte maturation [104,105]. Since then, multiple studies have assessed the development of OHSS in women who receive GnRH agonist trigger compared with hCG trigger for final oocyte maturation, resulting in a significant reduction in the development of OHSS. An analysis of 17 RCTs that assessed GnRH agonist compared with hCG trigger found that the agonist resulted in a lower incidence of OHSS in fresh autologous cycles (OR 0.15, 95% CI: 0.05–0.47), as well as in donor-recipient cycles (OR 0.05, 95% CI: 0.01–0.28) [120] compared with hCG. These studies also reported, however, that agonist trigger was associated with a lower live-birth rate (OR 0.47, 95% CI: 0.31–0.70) in fresh autologous cycles due to a luteal phase insufficiency [120]. That said, several strategies have been implemented to overcome this problem when embryo replacement is performed in the same cycle. An alternative consists of inducing corpus luteum function with low-dose (1500 IU) hCG. When used in women at risk of OHSS, a single 1500 IU hCG injection administered the day of oocyte retrieval is associated with no OHSS and sustainable pregnancy rates [121]. If the patient is not at risk, a second 1500 IU hCG injection can be administered 5 days later. However, some cases of late-onset OHSS have been described, highlighting the

fact that embryo transfer is a major driver of OHSS regardless of the number of follicles developed [121]. Therefore, the “freeze-all” policy should always be considered as a viable option in these cases. It is advisable to measure serum LH on the day of triggering, as the chance of having a suboptimal response to GnRH agonist triggering 12 h after administration (LH < 15 UI/mL) is as high as 25% if serum LH is undetectable, compared with 5% in the general population [122]. The suboptimal response is particularly evident in women with anovulatory irregular cycles and when taking oral contraceptives for a long period of time. Another option for overcoming a potential luteal phase defect is supplementing the luteal phase with both estrogens and progesterone according to the standard steroid replacement protocols. We usually employ 6 mg/day oral estradiol valerate and 800 mg/day vaginal micronized progesterone as in the oocyte donation cycles [123]. Using this luteal phase support, pregnancy rates are similar regardless of the triggering done with hCG or GnRH agonists [124]. Lowering or Withholding hCG Another alternative that has been tried in women with high risk of OHSS is to reduce the dose of hCG used to trigger ovulation, which has traditionally been 5000–10,000 IU of urinary hCG, and more recently 2500 IU recombinant hCG. Different studies have compared 5000 IU vs 10,000 IU, as well as 4000 IU vs 6000 IU, without any beneficial effect on the incidence of OHSS [125,126]. The only way to prevent OHSS if ovulation triggering must be performed with hCG (e.g., in long-GnRH agonist cycles) is to withhold hCG administration and avoid sexual intercourse. This decision is sometimes difficult as patients have already invested substantial time, money, and energy in the process, but many times it is worth canceling the cycle to avoid putting the patient at high risk. Kisspeptin (Kp) Kisspeptins are a group of hypothalamic peptides that are essential for normal human fertility [127]. Within the hypothalamus, Kp is released from kisspeptin-neurokinin B-dynorphin (KNDy) which directly projects to GnRH neurons, subsequently activating the secretion of LH and FSH [128,129]. Peripheral injection of Kp has been shown to potently stimulate gonadotropin secretion in humans through a GnRH-dependent mechanism [130–132]. Initial proof-of-concept studies have shown that Kp-54 administration has an effect on egg maturation in women undergoing in vitro fertilization (IVF). Following superovulation with recombinant FSH, women were administered a single subcutaneous injection of different doses of Kp-54 to induce an LH surge and egg maturation. Oocyte maturation was observed in response to each tested dose

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of Kp-54, and the mean number of mature eggs per patient generally increased in a dose-dependent manner [133]. In a subsequent study, the authors challenged the concept that Kp-54 would effectively trigger oocyte maturation and also have a low risk of inducing OHSS [134]. Following a standard recombinant FSH/GnRH antagonist protocol, patients were randomly assigned to receive a single injection of Kp-54 to trigger oocyte maturation using different doses. Women were routinely screened for the development of OHSS. Oocyte maturation occurred in 95% of women, with the highest oocyte yield following 12.8 nmol/kg Kp-54. No woman developed moderate, severe, or critical OHSS [134]. Kisspeptin is not yet commercially available.

After Egg Retrieval Oocyte/Embryo Cryopreservation One of the main advances that has changed clinical practice in ART has been the substantial and reproducible improvement in oocyte and embryo vitrification in terms of survival and overall health of newborn infants [135]. As a result, many cycles are associated with freezing all available oocytes or embryos. The policy is referred to as a “freeze-all policy” or “cycle segmentation.” The best OHSS preventive measure is oocyte/embryo cryopreservation, although there is not much literature on this method [136,137]. The reason is clear: avoiding pregnancy is the best way to avoid late-onset OHSS, the most dangerous form of the syndrome. Oocytes might be frozen or, alternatively, some can be fertilized and cryopreserved as embryos. Cryopreservation has also been shown to be more effective than coasting in earlyonset OHSS prevention [138]. Thus, a widely employed strategy in women at risk of OHSS is to use a combination of GnRH agonist triggering and cryopreservation of oocytes (the so-called freeze-all strategy). But even in the absence of embryo replacement, some cases of OHSS have been described in high-risk populations [139–141]. Therefore, it might be worth exploring other initiatives during the luteal phase in order to prevent OHSS in the absence of embryo replacement. Dopamine Agonists As previously described, an increased vascular permeability of the ovarian capillaries caused by ovarian hypersecretion of VEGF and increased VEGFR-2 expression is the main pathophysiology of OHSS [11,32,41]. We introduced the concept of targeting VEGF by administering dopamine agonists in the murine model [41]. Our first study in humans employed cabergoline 0.5 mg/day from the day of hCG for 8 days. The study was performed in oocyte donors and showed a significant reduction of moderate OHSS in the dopamine agonist group

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compared with controls [42]. A subsequent study included women undergoing a full IVF treatment including embryo replacement [43]. In this study, women were treated with different doses of the dopamine agonist quinagolide (0, 50, 100, or 200 mg/day until the day of pregnancy test). Quinagolide reduced ascites formation and moderate-severe OHSS in women who did not get pregnant, but it was ineffective in women who became pregnant. This may be either because the amount of trophoblast-derived hCG was too elevated to be compensated by the doses of quinagolide administered, or because pregnancy stimulates other pathways that also affect vascular permeability irrespective of VEGF blockage by a dopamine agonist. Subsequent analysis of the literature confirmed that administration of dopamine agonists reduced the incidence of OHSS compared with no treatment (RR 0.38, CI: 0.29–0.51) without affecting pregnancy rates (RR 1.02, 95% CI: 0.78–1.34) [142]. It was also more effective in early-onset than late-onset prevention, showing the difficulty of targeting pregnant women with this therapy [143]. Whether a more aggressive intravenous administration of dopamine in women developing severe/critical OHSS will work has not been studied, but there are some reports in which this strategy has successfully rescued renal output in compromised cases [144,145]. Albumin The rationale for the use of albumin in women at risk of OHSS is its ability to increase plasma oncotic pressure and counteract the permeability effect of angiotensin II. Moreover, albumin may also bind to vasoactive substances such as VEGF. However, the data evaluating the efficacy of albumin in the prevention of OHSS are contradictory. Initial early RCTs demonstrated that 20% human albumin administered at oocyte retrieval decreased the incidence of moderate-to-severe OHSS compared with no treatment [146]. In an RCT involving almost 500 patients per arm, patients received either 40 g of 20% human albumin intravenously just after pick up for 30 min, or no treatment [147]. The incidence of moderate (7.1% vs 6.7%) and severe OHSS (5.0% vs 4.7%) was similar between treatment and nontreatment groups, as well as the patients that needed culdocentesis. There was also no difference in blood parameters and hypercoagulability state. It was concluded that albumin was not useful for preventing hCG-induced OHSS when administered after oocyte retrieval [147]. Subsequent systematic reviews also concluded that albumin does not prevent OHSS [148,149]. Moreover, other studies have compared the use of human albumin to other methods of reducing OHSS risk such as hydroxyethyl starch solution (HES), coasting, or placebo and found that human albumin does not offer a

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significant benefit [150,151]. It is also worth noting that albumin is a blood-derived product, and can lead to allergic and anaphylactic reactions, and potentially to the transmission of viral or unidentified diseases. Calcium The rationale for calcium (Ca) administration has been to inhibit cAMP-stimulated renin secretion, which decreases angiotensin II synthesis and in turn might impair VEGF production. Thus, intravenous Ca infusion (10 mL of 10% Ca gluconate in 200 mL normal saline) on the day of oocyte retrieval and on days 1, 2, and 3 after oocyte retrieval, has been tested in an RCT of women at risk for OHSS. Results showed a significant reduction in the incidence of moderate and severe OHSS compared with saline (23% vs 7%) [152]. Ca seems to be as effective as dopamine agonists [153]. Letrozole Letrozole is a potent aromatase inhibitor that also effectively lowers serum estradiol levels in the luteal phase [154]. Thus, there is a strong rationale for employing letrozole as a luteolytic agent, and in fact a pilot study successfully employed letrozole to prevent OHSS [155]. An RCT has compared the use of letrozole (5 mg/day for 5 days starting the day of oocyte retrieval) with aspirin (100 mg/day for 5 days after pickup) in women at high risk of developing OHSS who received hCG to trigger ovulation and all the embryos were frozen. The incidence of ascites formation and overall moderate early-onset OHSS was reduced in the letrozole group compared with the aspirin-treated group 7 days after hCG (19.6% vs 35.3%, respectively), and the luteal phase was shortened by almost 3 days [156]. The study intended to show that letrozole might decrease serum VEGF levels as had been described [157], but in fact women treated with letrozole had higher serum VEGF levels than those treated with aspirin. It was inferred that letrozole might have luteolytic actions through different unknown mechanisms, which may be related to its androgenic effects on granulosa cells [156]. GnRH Analogs in the Luteal Phase GnRH analogs can be employed in two different directions during the luteal phase. One approach to induce luteolysis in women in whom a freeze-all policy has been applied is to continue GnRH antagonist administration during the luteal phase. In one report, 2 days of GnRH agonist administration plus dopamine agonists for 7 days prevented OHSS in a short series of patients [158]. Although no RCTs have been published, the potent luteolytic action of GnRH antagonists suggests that it may help to prevent OHSS after a freeze-all strategy. A different strategy is to perform embryo replacement in cycles at high risk of OHSS in which triggering has

been induced with an agonist. The luteal phase has been supported in a study including 46 patients with 200 mcgs for 2/day during the luteal phase until the day of the pregnancy test, where it was discontinued regardless of the results. No OHSS was observed, while the pregnancy rate was 52% [159]. Combination of Treatments Based on the information reported above, different steps should be considered to prevent OHSS. The first step is to always perform COS with GnRH antagonists protocols, unless the patient systematically develops an asynchronic follicular growth that cannot be managed with steroids or oral contraceptives [160,161]. This is especially important in women with elevated serum AMH, a high AFC, or for those who have experienced OHSS in previous cycles. GnRH antagonists protocols are associated with similar outcomes as GnRH agonists protocols [106], less gonadotropin consumption, and lower OHSS incidence, even if hCG is employed to trigger maturation [107]. In the course of COS, the clinician will determine the number of follicles developed and serum estradiol levels to move to step 2. Special attention should be paid to those individuals producing >19 follicles or serum estradiol >3500 pMol/L, but to be on the safe side we would recommend continuing to step 2 when >14 follicles have developed and serum estradiol >2500 pMol/L. In those patients, step 2 will consist of administering a GnRH antagonist to trigger ovulation and counting the number of oocytes retrieved. If >15 oocytes are obtained, step 3 will consist of freezing all the oocytes/embryos and perhaps the administration of either dopamine agonists, letrozole, GnRH antagonist, or a combination of those during the luteal phase, depending of the number of oocytes finally displayed and the signs and symptoms of OHSS. If the number of eggs collected is 45% WBC > 25,000L Creatinine > 1.6 mg/dL Oral analgesia inadequate for abdominal pain Tense ascites, hypotension Other severe complications or symptoms Yes Hospitalization: Transfer to centre with OHSS experience Inpatient management for severe OHSS Intensive care unit for critical OHSS Evaluation and monitoring: Daily weights and abdominal circumference measurements Laboratory testing: CBC, electrolytes, BUN, creatinine, liver enzymes, serum hCG (to determine if patient has conceived) TVUS as needed to monitor ovarian size and ascites Chest radiograph an echocardiogram if pleural and/or pericardial effusion suspected Invasive monitoring central venous pressure Management: Supportive care IV hydration Culdocentesis for removal of tense ascites causing significant pain or respiratory compromise Prophylactic anticoagulation in ALL patients with OHSS requiring hospitalization Thoracentesis for symptomatic pleural effusions: no data Pain management: acetaminophen, oral or parenteral opiates if needed; no NSAIDs or antiplatelet drugs Antiemetics if needed Management of other complications: multidisciplinary team (Internal medicine, admitting OB-GYN, including subspecialists, critical care if needed)

Resolution

No Continue out patient management and add: Prophylaxis for thromboembolism if two of three are present: age > 35 years, obesity, immobility, personal or family history of thrombosis or thrombophilia, pregnancy

Outpatient management for total of approximately two weeks or until menses (if not pregnant)

If nor pregnant, resolution over 10 – 14 days If pregnant, delayed resolution

Discharge when stable and monitor as outpatient

FIG. 3

Management of OHSS. (Reproduced with permission from Ref. [162], http://www.uptodate.com.)

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considered conservative and directed to treat symptoms. Most people can be managed as outpatients, but some women with severe or critical OHSS will require hospitalization, possibly in the ICU [162].

Mild OHSS Most OHSS cases are mild or moderate and can be managed on an outpatient basis with the goal of relieving symptoms because they are self-limited. For mild OHSS, analgesics and avoidance of heavy physical activity are usually enough. Nonsteroidal antiinflammatory drugs are contraindicated because they can compromise renal function, and therefore acetaminophen is usually administered. Patients should be instructed to call if any signs or symptoms become worse, especially with oliguria, abdominal distention, shortness of breath, or weight gain (>1 kg/day) because progression to moderate or severe forms can be expected in case of pregnancy (Fig. 3) [162].

by an increase in urine output [164]. Aspiration of >4 L of fluid at once is not recommended. It has been reported than >90% of patients will benefit from culdocentesis, and that a mean of 3.4 procedures are necessary until the syndrome is resolved [60,72]. The second aspect to be consider is replacement of at least part of the fluid removed. Culdocentesis must be always accompanied by intravenous hydration with isotonic crystalloid solutions (e.g., normal saline, Ringer’s lactate). The third important measure is thromboembolism prophylaxis. It should be considered in hospitalized patients and for women with OHSS being managed as outpatients with two of the following risk factors: age >35 years, obesity, immobility, personal or family history of thrombosis, thrombophilias, and pregnancy. We employ prophylactic low molecular weight heparin 20 mg subcutaneously every 12 h, or heparin 5000 units subcutaneously every 12 h [165].

Moderate OHSS

Severe and Critical OHSS

For women with moderate OHSS, recommendations include oral fluid intake of 1–2 L/day (Fig. 3) [162]. Diuretics are contraindicated because they can further decrease intravascular volume. It is important to instruct patients to perform daily recordings of weight, abdominal circumference, and urinary output. Regarding physical activity, they can ambulate but bed rest is sometimes necessary. Avoid other physical activity and especially sexual intercourse. Patients should be monitored in the clinic every 48–72 h, although any worsening should be reported and the patient should go to the clinic, as this often occurs when women become pregnant. These visits should control vital signs and perform a physical examination, an ultrasound to evaluate ascites, and laboratory testing of complete blood count, electrolytes, creatinine, serum albumin, and liver enzymes. In addition, three important measures should be considered. The first relates to reducing the volume of liquid accumulated in the abdomen. Although transabdominal paracentesis is reported to be successful [163], most centers use transvaginal aspiration of the ascitic fluid from the cul-de-sac using ultrasound to provide symptomatic relief. Even on an outpatient basis, culdocentesis is often performed in women with tense ascites, orthopnea, rapid increase of abdominal fluid, or any other sign that may indicate worsening [162]. Antibiotic coverage should also be used to avoid infections. The volume of fluid to be removed is a matter of controversy. In general, after aspiration of 500 mL of fluid, patients report resolution of abdominal discomfort. Aspiration of 2000 mL of ascites results in intraabdominal pressure reduction and renal artery resistance, followed

Hospitalization is mandatory in women with severe OHSS and any of the following criteria: a hematocrit >55%, leukocytes >25,000/L, and creatinine >1.6 mg/ dL. Also, women with severe abdominal pain, intractable vomiting, severe oliguria/anuria, tense ascites, dyspnea or tachypnea, hypotension, dizziness or syncope, severe electrolyte imbalance, or abnormal liver function tests must also be hospitalized (Fig. 3) [162]. Management of severe and critical OHSS is complicated, and therefore patients should be hospitalized in units with experience treating this condition. Alternatively, in the benefit of the patient or because of the severity of the syndrome, direct hospitalization in an ICU is strongly advised. Treatment will consist of supportive care, monitoring, and prevention and treatment of complications. The first goal is to maintain intravascular blood volume. Although isotonic crystalloids are typically used, some clinicians employ intravenous albumin due to its osmotic properties in order to shift the direction of fluid from the extravascular to the intravascular space. It has been shown, however, that intravenous albumin provides no additional benefit when compared with crystalloid solutions. The only indication for using albumin is when there is concomitant oligoanuria. In this case, it is advised to administer intravenous albumin (25% in 250 mL) and subsequently furosemide (20mg) in order to augment urine output. Also, intravenous dopamine (2–3 μg/kg/min) can be employed to force urinary output and repeated every 8 h, but these measures are performed by the ICU staff [144,145]. Critical OHSS cases should be managed in an ICU because they can be complicated with massive

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REFERENCES

hydrothorax, pericardial effusion, arterial thrombosis, pulmonary embolism, sepsis, acute renal failure, ARDS, and DIC, all of which need to be managed by specialized staff [165]. Oophorectomy is never an option in life-threatening conditions, but pregnancy interruption might be necessary in uncontrolled cases after unsuccessful application of available therapeutic approaches. As already mentioned, OHSS is self-limited and increased ovarian vascular permeability regresses spontaneously in 10–14 days. The fluid in the third space slowly reenters the intravascular space, hemoconcentration reverses, and natural diuresis ensues. As a result, hematocrit is normalized, ultrasound shows no signs of ascites, and in general clinical symptoms alleviate. If pregnancy occurs, resolution may take longer [166]. Some studies suggest that pregnancies complicated with OHSS have a higher rate of miscarriage, gestational diabetes, and pregnancy-associated hypertension [167,168], but these complications have not been confirmed in other studies [169].

CONCLUSIONS OHSS is a complication of infertility treatments that is totally avoidable today. The syndrome consists of a series of signs and symptoms which are the consequence of increased ovarian capillary permeability after hCG administration. Pain and discomfort due to enlarged ovaries, ascites, hypercoagulability, and reduced renal function are common findings. OHSS is usually mild, but can be complicated seriously and become a lifethreatening condition, especially in women who become pregnant. Some phenotypic characteristics, such as race, PCOS, and biomarkers of ovarian reserve (AFC and AMH levels) predict the risk of developing OHSS in infertile patients. Special protocols for ovarian stimulation using low-dose gonadotropins, GnRH antagonists to prevent endogenous LH surges, and GnRH agonists to trigger ovulation, should be employed to minimize risk. Moreover, cryopreservation of oocytes/embryos and deferring pregnancy for a subsequent cycle is a very important step in the prevention of OHSS. When OHSS develops, treatment consists of the management of symptoms and monitoring of vital signs, including culdocentesis of ascitic fluid, intravenous liquid replacement, antithrombotic measures, and pain reduction. These patients should be managed in specialized units to ensure the best possible outcomes.

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[161] Garcia-Velasco JA, Bermejo A, Ruiz F, Martinez-Salazar J, Requena A, Pellicer A. Cycle scheduling with oral contraceptive pills in the GnRH antagonist protocol vs the long protocol: a randomized, controlled trial. Fertil Steril 2011;96(3):590–3. [162] Busso CE, Soares SR, Pellicer A. Management of ovarian hyperstimulation syndrome. In: UpToDate, Post TW (Ed), UpToDate, Waltham, MA (Accessed on June 19, 2017.) [163] Levin I, Almog B, Avni A, et al. Effect of paracentesis of ascitic fluids on urinary output and blood indices in patients with severe ovarian hyperstimulation syndrome. Fertil Steril 2002;77(5):986–8. [164] Maslovitz S, Jaffa A, Eytan O, et al. Renal blood flow alteration after paracentesis in women with ovarian hyperstimulation. Obstet Gynecol 2004;104(2):321–6. [165] Busso CE, GarciaVelasco JA, Gomez R, et al. Ovarian hyperstimulation syndrome. In: Rizk B, GarciaVelasco JA, Sallam HM, Makrigiannakis A, editors. Infertility and assisted reproduction. Cambridge, UK: Cambridge University Press; 2008. p. 243. [166] Nouri K, Tempfer CB, Lenart C, et al. Predictive factors for recovery time in patients suffering from severe OHSS. Reprod Biol Endocrinol 2014;12(7):59. [167] Abramov Y, Elchalal U, Schenker JG. Obstetric outcome of in vitro fertilized pregnancies complicated by severe ovarian hyperstimulation syndrome: a multicenter study. Fertil Steril 1998;70 (6):1070–6. [168] Mathur RS, Jenkins JM. Is ovarian hyperstimulation syndrome associated with a poor obstetric outcome? BJOG 2000; 107(8):943–6. [169] Wiser A, Levron J, Kreizer D, et al. Outcome of pregnancies complicated by severe ovarian hyperstimulation syndrome (OHSS): a follow-up beyond the second trimester. Hum Reprod 2005; 20(4):910–4.

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23 The Role of GnRH Agonist Triggering in GnRH Antagonist-Based Ovarian Stimulation Protocols Shahar Kol, Ofer Fainaru Abbreviations CL E2 EFS ET FSH GnRH GnRHa GnSAF hCG ICSI IM IU IVF LH LPS OHSS P r-hCG u-hCG

corpus luteum, or corpora lutea for plural estradiol empty follicle syndrome embryo transfer follicle-stimulating hormone gonadotropin-releasing hormone gonadotropin-releasing hormone agonist gonadotrophin surge attenuating factor human chorionic gonadotropin intracytoplasmic sperm injection intramuscular international units in vitro fertilization luteinizing hormone luteal-phase support ovarian hyperstimulation syndrome progesterone recombinant human hCG urinary-derived hCG

INTRODUCTION The first successful human in vitro fertilization (IVF) treatment was reported in 1978 [1]. It was achieved by harvesting an oocyte by laparoscopy in a natural cycle. This approach was associated with very low chances for live birth, and was soon replaced by strategies that are more efficient. Final oocyte maturation and the release of a viable, fertilizable oocyte from the ovarian follicle are the hallmark of natural mid-cycle luteinizing hormone (LH) and follicle-stimulating hormone (FSH) surges. For in vivo fertilization, a mature oocyte is extruded from the ovulating follicle to the adjacent fallopian tube, where fertilization takes place. For in vitro fertilization, mature oocytes are retrieved before follicular rupture. Since oocytes are arrested at the prophase of

The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00023-6

the first meiotic division, the trigger (either spontaneous or exogenously administered) initiates a cascade of events that results in meiosis completion, luteinization of the follicular wall, and ovulation. In clinical practice, choosing the right ovulation trigger must take into account the follicular phase preceding the trigger, the hormonal environment at the time of trigger, and the implications of the trigger on the following luteal phase. All these considerations are pivotal to support implantation of an embryo in the uterus, establishing a viable pregnancy, and at the same time to minimize risks, particularly, ovarian hyperstimulation syndrome (OHSS). For practical purposes, human chorionic gonadotropin (hCG) was chosen as the triggering agent, when controlled hyperstimulation was introduced to IVF treatment. In parallel, GnRH agonists (GnRHa) became available, and soon dominated the IVF arena, given their capacity to induce pituitary downregulation, and to minimize the risk of premature luteinization during ovarian stimulation [2]. In the late 1990s of the 20th century, GnRH antagonists first became available, opening the option to trigger ovulation with GnRHa. This practice soon gained popularity. Indeed, in 2013, >36% of IVF cycles in Europe were triggered with GnRHa [3]. The purpose of this chapter is to review the use of GnRHa for the purpose of final oocyte maturation, with particular emphasis on the luteal phase that follows.

THE MID-CYCLE OVULATION TRIGGER IN NORMAL PHYSIOLOGY The purpose of the ovarian cycle is to produce a single, healthy oocyte that will be fertilized leading to a viable

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pregnancy. During the follicular phase, the dominant follicle (with its components: an oocyte arrested in the prophase of the first meiotic division, granulosa and theca cells, basement membrane) acquires the properties and maturation stage required for ovulation. The cue for ovulation is controlled by the pituitary, producing a biochemical trigger (LH and FSH surges), when the dominant follicle is ready. Hoff and Quigley [4] described the hormonal events that characterize the surges. The onset of LH and FSH surges occurs abruptly (LH doubled within 2 h). The surges are temporarily associated with peak estradiol (E2) levels and occur 12 h after the initiation of a rapid rise of progesterone (P). The mean duration of the surge is 48 h, with a rapidly ascending limb (doubling time, 5.2 h) lasting 14 h, accompanied by a rapid decline of E2 and a continued rise of P. A peak plateau of LH and FSH levels lasting for 14 h and a transient leveling of P follow the surge. The longer descending limb (halftime, 9.6 h), lasting for 20 h, is associated with a second rapid rise of P, beginning 36 h after surge onset or 12 h before termination of the surge. If we break down oocyte maturation events during the surge, it appears that by 18 h after surge onset, resumption of meiosis has occurred, while 28–36 h into the surge, metaphase II oocytes can be harvested [5]. However, as discussed later, shorter surges, like those triggered with GnRHa, are also associated with recovery of mature oocytes. The threshold amplitude of LH required for human oocyte maturation and ovulation is not known, although a level of 100 IU/L is considered normal. FSH surge levels tend to reach an amplitude of 25–30 IU/L [6]. Notably, ovulation physiology changes with age. Older women present with larger initial follicle diameters than younger women and ovulate at a smaller mean follicle diameter. Follicle growth begins earlier in the cycle of older women, but growth progresses more slowly. Hence, ovulation may occur at an earlier stage of growth in association with reproductive aging [7].

hCG-BASED TRIGGERING IN OVARIAN STIMULATION CYCLES The hCG molecule is routinely used for triggering ovulation in the clinical setting because of its ability to activate the ovarian LH receptor, and since it was the first triggering agent that was clinically available. hCG, produced by trophectoderm of the implanting embryo, is the first biochemical signal of a new pregnancy, and can be detected in maternal serum 8–10 days after ovulation in a fertile cycle, which coincides with trophoblast formation and implantation of the blastocyst in the endometrium. hCG serves as a signal to the ovary to maintain corpus luteum function, which would regress, if

not rescued by hCG. Corpus luteum function is vital for maintaining receptive endometrium. hCG is a heavily glycated glycoprotein with a molecular weight of about 36,700 Da, of which 30% is sugar residues. It is composed of α and β subunits, which are noncovalently linked. The β subunit confers specific activity to the hormone. Much homology exists between hCG and LH, especially with respect to the first 121 amino acids of the β subunits of both hormones, which have about 80% homology. This homology enables hCG to serve as a strong ligand for the LH receptor in the follicular unit, hence its widespread clinical use. hCG has a 24-amino acid extension on the carboxyl-terminal end that is lacking in the LH β subunit. This C-terminal peptide of hCG-β markedly increases the half-life of hCG in vivo [8]. Extended half-life, LH homology and a relatively easy manufacturing process made hCG an excellent molecule to be used in triggering ovulation within the framework of assisted reproductive technology treatments. The pharmaceutical industry provides us now with urinaryderived hCG (u-hCG), or recombinant human hCG (r-hCG), that are routinely used to trigger and time ovulation. Typically, one bolus of hCG (5000–10,000 units in the case of urinary derived products, or 250 μg in the case of a recombinant molecule) is administered 36 h before oocyte retrieval in IVF cycles. The optimal dose of urinary hCG that can trigger final oocyte maturation, leading to higher IVF success rate, without increasing the risk of OHSS is not known, and is probably patient specific [9]. A 190-patient randomized controlled study was conducted in order to compare the effectiveness of 250 μg r-hCG and 5000 IU u-hCG. The conclusion was that r-hCG is effective in inducing ovulation trigger, and is associated with more mature oocytes, higher P concentration, and improved patient tolerance compared with u-hCG [10]. When two doses of r-hCG (250 and 500 μg) were compared in a randomized control trial, the numbers of two pronuclei fertilized oocytes on day 1 after oocyte retrieval, or cleaved embryos on the day of embryo transfer (ET), were significantly higher with 500 μg of recombinant hCG than with the lower dose. However, the incidence of adverse events also tended to be higher with the higher dosage [11]. Unlike the LH/FSH surge, that terminates 48 h after its onset, hCG-mediated LH activity extends long into the luteal phase. This supraphysiologic LH activity overstimulates the corpora lutea (CL), leading to high serum E2 and P levels, that in turn decrease endogenous LH secretion [12]. hCG administered for final oocyte maturation supports the luteal phase for a maximum of 8 days; however, since endogenous LH is low, a luteal-phase defect ensues, which makes luteal-phase support (LPS) mandatory in order to maintain a receptive endometrium [13]. Studies in humans and primates have demonstrated that

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THE BENEFIT OF DUAL (LH AND FSH) TRIGGERING

the CL requires a consistent LH stimulus in order to function [14]. LH support during the luteal phase is entirely responsible for maintaining normal steroidogenic activity of the CL [15]. As a result, withdrawal of LH causes premature luteolysis [16]. LH secretion pattern in the luteal phase (baseline level, pulse frequency, and amplitude) is described elsewhere [17]. In summary: for ovulation triggering, clinicians have what is easy to produce and use clinically (urinary or recombinant hCG), however, with a price to pay for significant deviation from physiology, particularly, in the luteal phase.

THE RATIONAL FOR GnRHa TRIGGERING The dynamics of the GnRHa-driven triggering were described by Itskovitz et al. [18]. A rapid LH ascending limb was demonstrated, reaching maximal LH levels within 4 h, followed by a longer descending limb of 16 h. The LH amplitude was comparable with its physiologic counterpart; however, it was significantly shorter. This significant deviation from physiology apparently does not hamper GnRHa trigger ability to produce mature oocytes for IVF as was repeatedly shown. The parallel FSH surge described by Gonen et al. [19] is also comparable in amplitude but shorter in duration compared with normal physiology (Fig. 1). It is important to emphasize that these two studies were conducted before the GnRH antagonist era. Once GnRH antagonists were introduced, it became necessary to demonstrate that the antagonist molecule can be displaced from pituitary receptors by the agonist molecule. Indeed, Fauser et al. [20] showed in a randomized, multicenter study, that GnRHa could adequately trigger ovulation in a GnRH antagonist-based ovarian stimulation protocol. Both GnRHa preparations used in the study (triptorelin and leuprorelin) elicited comparable LH surges. The time and amplitude patterns of LH and FSH surges in the study were identical to those reported without GnRH antagonist cotreatment [18], suggesting that the agonists completely displace the antagonist from the pituitary receptors (Fig. 2). E2 and P secretion patterns GnRHa LH

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during the first 36 h post-GnRHa triggering were identical to those observed following hCG triggering. Moreover, intramuscular P injection-based luteal support following the GnRHa trigger secured clinical outcome comparable to those seen following hCG trigger (control group).

THE BENEFIT OF DUAL (LH AND FSH) TRIGGERING Although the role of the FSH mid-cycle surge is not fully explored, it is known to promote LH receptor formation in luteinizing granulosa cells, nuclear maturation (i.e., resumption of meiosis), and cumulus expansion [21–23]. Since hCG offers LH activity only, while in the natural cycle it is accompanied by an FSH surge, investigators explored the possibility of adding FSH as a “dual surge” in an effort to mimic physiology more closely. Lamb et al. [24] added 450 IU of FSH to the routine 10,000 IU u-hCG trigger dose in a randomized double blind placebo controlled study. The dose of FSH was chosen to imitate the 1:4 ratio (FSH:LH) observed in the natural cycle. Fertilization rate was significantly improved in the treatment (FSH + hCG) arm compared with control (63% vs 55%), as was the likelihood of oocyte recovery (70% vs 57%). Nevertheless, there was no significant difference in clinical pregnancy rate (56.8% vs 46.2%) or ongoing/live birth rate (51.6% vs 43.0%). Others have studied GnRHa and hCG “dual triggering,” making use of the GnRHa ability to induce an FSH surge in addition to the LH surge [19]. Shapiro et al. [25] supplemented a GnRHa trigger with a relatively small dose of hCG. A series of 45 patients at high risk for OHSS received injections of leuprolide acetate (4 mg) and hCG (1000–2500 IU). The hCG dose varied according to the patient weight and OHSS risk factors. No patient developed OHSS; and a 53% ongoing pregnancy rate was achieved. The authors’ conclusion of this “proofof-concept” study was that concomitant administration of GnRHa and hCG for final oocyte maturation appears to be effective and safe in high responders in terms of the ability to achieve ongoing pregnancies and at the same time reducing the risk for OHSS. Similar results were published for patients with low proportion of mature-MII oocytes [26]. Surprisingly, a bolus of FSH can replace hCG as a trigger as described in a case report by Bianchi et al. [27]. Inadvertently, a patient injected herself with 2010 units of recombinant FSH instead of hCG 36 h before retrieval. Out of the nine oocytes retrieved, eight were metaphase II. Three oocytes were injected with sperm, and developed into good quality embryos, although failed to achieve pregnancy. This intriguing case report suggests

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FIG. 2 Serum concentrations of LH (hCG), FSH, E2, and P (mean SEM) during triggering of final stages of oocyte maturation (0, 4, 8, 12, and 24 h) with two GnRH agonist (triptorelin and leuprorelin) or hCG after ovarian hyperstimulation for IVF and during the subsequent luteal phase [day of oocyte pick-up (OPU), embryo transfer (ET), and 1 and 2 weeks after ET]. From Fauser B, de Jong D, Olivennes F, Wramsby H, Tay C, Itskovitz-Eldor J, et al. Endocrine profiles after triggering of final oocyte maturation with GnRH agonist after cotreatment with the GnRH antagonist ganirelix during ovarian hyperstimulation for in vitro fertilization. J Clin Endocrinol Metab 2002;87(2):709–715, by permission.

that an FSH surge alone may be sufficient to promote final oocyte maturation, similar to hCG. It also hints to the fact that the natural FSH surge is not physiologically redundant. In summary, although the body of evidence is not large enough to draw firm conclusions, there seems to be an advantage in adding FSH (as FSH itself, or GnRHa-driven) to hCG triggering of ovulation.

GnRHa TRIGGERING AFTER OVARIAN STIMULATION WITH GnRH ANTAGONIST COTREATMENT Are all GnRHa equal? Parneix et al. [28] explored different types, doses and modes of administering GnRHa in 123 women, although no significant differences were found. Others have used a wide range of GnRHa preparations: triptorelin 0.2 mg [29–31], buserelin 0.5 mg [32], leuprolide acetate 1 mg [33], or leuprolide acetate 1.5 mg [34]. A formal pharma-initiated “dose finding” study was never conducted, probably since the use of GnRHa as ovulation trigger is not endorsed by the manufacturers and is still “off-label,” in spite of its wide use. Vuong et al. [35] showed no significant differences between triptorelin doses of 0.2, 0.3, and 0.4 mg used

for ovulation trigger in oocyte donors (n ¼ 165) with regard to the number of mature oocytes and top-quality embryos achieved.

Number of Mature Oocytes Retrieved and Embryo Quality In a prospective randomized study, Humaidan et al. reported significantly more mature oocytes retrieved following GnRHa trigger compared with hCG trigger [32]. Similar results were reported by Shapiro et al. [25] in an oocyte donation model: donors triggered with GnRHa produced a mean of 28.8 oocytes, compared with 21.7 oocytes retrieved after hCG trigger (P ¼ .003). In a randomized control study, Krishna et al. [36] showed that patients triggered with GnRHa compared with patients triggered with HCG had more oocytes (23.5 vs 20.8, P ¼ .006), more metaphase II oocytes (19 vs 14, P < .001), higher fertilization rates (86% vs 78%, P ¼ .001) and more top quality embryos (91% vs 74%, P ¼ .002). Empty follicles post-GnRHa triggering: in a large study, exploring the incidence of empty follicle syndrome (EFS) after GnRHa triggering vs hCG triggering in >3000 patients, Castillo et al. [37] showed that the incidence of EFS seems to be similar regardless of whether GnRHa or hCG triggering was used for final oocyte maturation.

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LUTEOLYSIS POST-GnRHa TRIGGERING

In our experience, EFS can occur if GnRHa is administered too soon after the last GnRH antagonist dose. We, therefore, recommend withholding this last dose or alternatively making sure that GnRHa is administered at least 8 h after the last GnRH antagonist injection.

GnRHa TRIGGERING FOR PREVENTING OHSS Pre-GnRH Antagonist Era The first report on the use of GnRHa for ovulation triggering in the context of OHSS prevention was published in 1988 as an abstract [38] followed by a full paper, underscoring the concept that a bolus of GnRHa is able to trigger an adequate mid-cycle LH/FSH surge, resulting in oocyte maturation and pregnancy. In addition, experience with eight patients with exaggerated response suggested that it might prevent clinical manifestations of OHSS [18]. As more experience was gained with GnRHa triggering, it became apparent that a randomized controlled study, comparing hCG and GnRHa triggers in high-risk patients, would be problematic to conduct. Therefore, a case-control study was published in an effort to describe the unparalleled strength of this modality in comparison with other strategies used to prevent OHSS [39]. At first, the IVF community was reluctant to adapt GnRHa triggering in the context of OHSS prevention, since up to the year 2000 GnRH agonists were used solely to prevent premature LH rise and luteinization by pituitary downregulation in most IVF cycles. Naturally, under these circumstances, GnRHa could not be used as a trigger. In addition, the reported experience of total elimination of OHSS was met with skepticism and disbelief. In order to allow GnRHa triggering, a GnRH analog-free protocol must be used, with an increased risk of cycle cancellation due to premature LH rise and luteinization. On the other hand, GnRHa triggering was utilized in “hyper responder” patients, in whom high serum concentrations of gonadotrophin surge attenuating factor (GnSAF) are found. GnSAF decreases the risk of premature luteinization by its action on endogenous GnRH secretion pattern [40,41]. Therefore, in IVF patients at high risk of OHSS, the risk of premature luteinization and cycle cancellation is actually low, even if GnRH agonists are not used for pituitary downregulation.

GnRH Antagonist Era During the last decade of the 20th century, intensive pharma efforts reached the target of producing GnRH antagonists with proven clinical activity and few side effects. As specified earlier, GnRH antagonists allowed

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the use of GnRHa for ovulation triggering. Since the last antagonist dose is given many hours before the GnRHa trigger dose, it seems plausible that agonist trigger under these circumstances can elicit adequate LH surge to secure final oocyte maturation. The first proof of concept study in the context of OHSS prevention was published in the year 2000 [42]. The first (and probably the last) randomized controlled study comparing OHSS incidence in “high-risk” patients triggered with a GnRHa or hCG was published in 2008 [33]. As expected, patients randomized to the GnRHa trigger did not develop OHSS at all, vs 30% of the patients randomized to the hCG arm. Multiple publications soon followed, as did meta-analyses confirming a reduced OHSS incidence following GnRHa triggering [43,44]. Few publications reported isolated cases of OHSS after GnRHa trigger cycles [45], yet, OHSS after GnRHa trigger and “freeze-all” strategy is very rare and probably relates to polymorphisms of the ovarian LH receptors.

LUTEOLYSIS POST-GnRHa TRIGGERING Although the efficacy of GnRHa triggering in preventing OHSS was evident, the mechanism is not clear. To shed more light on this aspect, luteal activity post-GnRHa triggering was examined [46]. Specifically, corpus luteum function was assessed by measuring luteal-phase levels of inhibin A and pro-αC, peptides that reflect luteal-phase activity. These peptides were measured in a small prospective randomized trial, after controlled ovarian hyperstimulation with FSH and GnRH antagonist. Following triggering of final oocyte maturation with either hCG (n ¼ 8) or GnRHa (n ¼ 8), blood was collected every 2–3 days during the luteal phase. Levels of inhibin A, pro-αC, E2, and P were significantly lower from day 4 to day 14 after triggering final oocyte maturation by GnRHa compared with hCG (Fig. 3). These results confirm that triggering final oocyte maturation with GnRHa instead of hCG in IVF cycles dramatically decreases luteal levels of inhibins, reflecting significant inhibition of corpus luteum function. The luteolysis pattern and timing after GnRHa triggering was demonstrated in four oocyte donors. On day 1 after triggering, circulating P concentrations are comparable, regardless of whether HCG or GnRHa were used for triggering. In contrast, P concentrations return to baseline 5 days after GnRHa triggering (without any luteal support), reflecting complete luteolysis by this day [47] (Fig. 4). As stated earlier, rare cases of OHSS post-GnRHa triggering prove that the luteolytic process post-GnRHa trigger is not universal [48]. Classical experiments [49,50], conducted decades ago, showed that luteal physiology in the natural cycle is based on pulsatile LH secretion. Specifically, Filicori

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et al. showed that mean LH pulse frequency declines from 15.2 pulses/24 h in the early to 8.4/24 h in the late luteal phase. A trend toward reduction in the amplitude of LH pulses was also observed [49]. In the context of ovarian stimulation for IVF and GnRHa triggering, corpora lutea function reflects endogenous LH secretion only, as is the case in a natural cycle. Recently, repeated blood sampling during early luteal-phase post-GnRHa triggering showed significant changes in LH secretion pattern [51]. Specifically, although pulsatile LH secretion continues, mean LH concentrations and LH pulse amplitude are significantly lower than those described in a natural cycle. During the first 48 h following oocyte retrieval P increases as seen following hCG trigger. The process of luteolysis starts very early in the luteal phase as indicated by decreasing P and E2 levels 2 days after ovulation (Fig. 5). Significant interpatient variability in terms of LH secretion pattern was noticed. Importantly, when hCG is given for luteal support, very high LH-like activity is reached, with no pulsatility at all, but rather a slow steady decline. It is tempting to speculate that the ovarian LH receptor responds to the ligand in two different patterns: in low ligand levels, pulsatility is necessary, but there is a threshold ligand concentration above which the receptor is activated regardless of pulsatility. This threshold level probably varies from patient to patient. In a patient with very low threshold, constant LH concentration (as seen after GnRHa triggering) may be enough to sustain the CL function. This may explain failure to achieve luteolysis post-GnRHa trigger in rare cases.

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FIG. 3 Luteal phase serum concentrations (mean SE) of inhibin A (A), pro-αC (B), P (C), and E2 (D) in two IVF protocols: GnRH antagonist for ovulation prevention and hCG (hCG group) or GnRHa (agonist group) for oocyte maturation triggering. Time is represented as days relative to oocyte maturation triggering (day 0). The changes in the levels of all four hormones in both groups were significant over time. P < .0001 (Friedman test). *P < .05; **P < .01; ***P < .001; and ****P < .0001. From Nevo O, alia Eldar-Geva T, Kol S, Itskovitz-Eldor J. Lower levels of inhibin A and pro-αC during the luteal phase after triggering oocyte maturation with a gonadotropin-releasing hormone agonist versus human chorionic gonadotropin. Fertil Steril. 2003;79(5):1123–1128, by permission.

As stated earlier, GnRHa triggering results in complete and quick luteolysis, as a rule. Luteolysis is achieved as early as 5 days post-oocyte retrieval, about 2 days before predicted implantation. As expected, nonsupplemented luteal phase following GnRHa triggering resulted in poor reproductive outcome [52], as one ongoing pregnancy was achieved in 15 patients so treated. LPS post-GnRHa trigger must take into account two considerations: first, since luteolysis occurs before embryo implantation, the required hormonal support is totally dependent on the supplement, unless an effort is made to rescue the corpora lutea. Second, there is a direct correlation between the E2 levels achieved during the follicular phase (ovarian stimulation) and the P levels required to support successful implantation. Standard LPS is routinely given post-hCG triggering in the form of vaginal P and oral E2. Indeed, early studies with standard LPS following GnRHa triggering yielded poor reproductive outcome [32,53].

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FIG. 4

Mean P values after GnRHa trigger. Blue line represents egg donors that did not receive any luteal support. LPS, luteal phase support. From Fatemi HM, Polyzos NP, van Vaerenbergh I, Bourgain C, Blockeel C, Alsbjerg B, et al. Early luteal phase endocrine profile is affected by the mode of triggering final oocyte maturation and the luteal phase support used in recombinant follicle-stimulating hormone–gonadotropin-releasing hormone antagonist in vitro fertilization cycles. Fertil Steril 2013;100(3):742-7.e1, by permission.

In a natural conception cycle, minimal mid-luteal P concentration to sustain pregnancy is in the range of 25–30 nmol/L [54]. Following ovarian stimulation, mid-luteal P must increase further, probably in a direct relation to higher maximal E2 levels achieved in the follicular phase, to allow for embryo implantation. In this case, minimal P level to sustain pregnancy is estimated to be in the range of 80–100 nmol/L [55]. Therefore, there is a prominent interest in securing a high luteal P concentration, especially during the implantation window. In order to secure high steroids (E2, and mainly P) levels in mid-luteal phase, two strategies were followed: • Administering high doses of P and E2. • Rescuing the corpora lutea to stimulate endogenous E2 and P production.

Intense LPS Engmann et al. [33] first described this approach in a randomized controlled study comparing hCG and GnRHa triggering in high-risk patients for developing OHSS. Patients allocated to the GnRHa trigger arm received E2 patches, intramuscular P and achieved high

implantation and pregnancy rates with no OHSS (Fig. 6). So far, this approach was not repeated in clinical trials, and in fact, the authors themselves subsequently deviated from this treatment plan and added low-dose hCG to the GnRHa at trigger time [56].

Modified Luteal Support With hCG GnRHa triggering opens the possibility to separate the required trigger properties from the LPS that follows. For triggering purposes, a large dose of hCG must be used (5000–10,000 IU), however, to sustain luteal function smaller doses suffice. Over the years, investigators presented different approaches for hCG-based luteal support post-GnRHa triggering in terms of timing and doses. • GnRHa triggering and early LPS with low-dose hCG combined with a standard LPS. In this strategy, a single bolus of hCG (1500 IU) is given shortly after oocyte retrieval, followed by standard LPS (oral E2 and vaginal P) (Fig. 7). This approach was assessed in a few trials, which proved it efficient in terms of reproductive outcome, with diminished risk of OHSS [57,58].

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Time (min) FIG. 5 Mean P and E2 levels after GnRHa trigger: Repeated measurements every 20 min started 48 h following oocyte retrieval (n ¼ 5). Based on data from Tannus S, Burke Y, McCartney CR, Kol S. GnRH-agonist triggering for final oocyte maturation in GnRH-antagonist IVF cycles induces decreased LH pulse rate and amplitude in early luteal phase: a possible luteolysis mechanism. Gynecol Endocrinol. 2017;33:741–745.

FIG. 6 GnRH antagonist-based ovarian stimulation followed by GnRHa triggering and intense luteal support with IM progesterone and transdermal E2 patches.

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FIG. 7 GnRH antagonist-based ovarian stimulation followed by GnRHa triggering and luteal support with 1500 IU of hCG on the day of oocyte retrieval, and standard luteal support with vaginal P and oral E2.

In a retrospective cohort study, using the same treatment scheme (including standard support), severe OHSS occurred in 6 of 23 high-risk patients (26%) [59]. • “Dual trigger”: GnRHa and a small hCG dose for triggering final oocyte maturation combined with intensive LPS. To boost luteal function, a single bolus of hCG 1000 IU is given together with GnRHa at trigger time. This strategy was compared with GnRHa trigger alone in a retrospective study [59]. The dual-trigger group had a significantly higher live birth rate (52.9% vs 30.9%), implantation rate (41.9% vs 22.1%), and clinical pregnancy rate (58.8% vs 36.8%) as compared with the GnRHa trigger group. One case of mild OHSS occurred in the dual-trigger group, and there were no cases of OHSS in the GnRHa trigger group. Of note, both groups received “intensive” luteal support with intramuscular P and transdermal E2 patches (Fig. 8). Mid-luteal P levels were identical in the two groups, suggesting that the small

hCG bolus at trigger time had no significant effect on endogenous luteal function. In a similar retrospective cohort study, dual trigger for final oocyte maturation using GnRHa and low-dose hCG was associated with a significantly increased risk of severe OHSS compared with GnRHa alone [60]. • GnRHa trigger and LPS with low-dose hCG 3 days after oocyte retrieval combined with intensive LPS. As shown earlier, adding hCG to the GnRHa trigger (either on trigger day or retrieval day) may expose patients to OHSS risk. Since early OHSS occurs 3–7 days posttriggering, more information regarding specific patient risk may be gathered by withholding the 1500 IU hCG luteal rescue bolus and re-evaluating these patients 3 days after oocyte retrieval for signs of early moderate OHSS. Only those patients with no signs of early OHSS can proceed with a bolus of HCG (1500 IU). This strategy was assessed in a small retrospective study [61]. No OHSS was reported. Of note, all patients

FIG. 8 GnRH antagonist-based ovarian stimulation followed by GnRHa triggering combined with 1000 IU of hCG (dual trigger) and intense luteal support with IM progesterone and transdermal E2 patches.

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FIG. 9 GnRH antagonist-based ovarian stimulation followed by GnRHa triggering and luteal support with 1500 IU of hCG 3 days after oocyte retrieval, and intense luteal support with intramuscular P, vaginal P, and oral E2.

received “intensive” luteal support with intramuscular P, vaginal P, and oral E2 (Fig. 9). Patients who received the 1500 IU hCG bolus had significantly higher mid-luteal P levels (P > 127 nmol/L). • GnRHa trigger and LPS with low-dose hCG after “luteal coasting,” without further progesterone support. The above strategies used fixed time points for the administration of low-dose hCG, i.e., either on the day of GnRHa trigger, on the day of oocyte retrieval, or 3 days after oocyte retrieval. To further individualize the LPS in OHSS high-risk patients, having a fresh transfer after GnRHa trigger, the same principle that holds for follicular phase coasting [62] might be valid during the luteal phase. In other words, monitoring P concentrations, and administering the hCG luteal-phase rescue bolus when P concentrations drop significantly. This strategy was described in a “case series” [63]. Since it was assumed that decreasing luteal P concentrations reflect luteolysis, luteal support with a single bolus of HCG

(1500 IU) was given when P concentration decreased below 30 nmol/L. No further luteal support was given (Fig. 10). Of the 21 patients so treated, 29% achieved clinical pregnancy. The only patient that developed early onset moderate OHSS had a mid-luteal P of 140 nmol/L. Importantly, in most patients in this series luteolysis began 2 days after oocyte retrieval; however, a few patients demonstrated delayed luteolysis (Fig. 11). • GnRHa trigger and LPS with daily microdose hCG without further progesterone support. hCG-based luteal rescue results in high mid-luteal P level, suggesting that further P supplementation is not needed. This concept was further explored in a proof-of-concept study [64]. It was shown that daily hCG dose (125 IU) supplementation in GnRHa triggered IVF cycles can replace standard support (Fig. 12). Interestingly, mid-luteal P was higher in the study groups (GnRHa for trigger, daily 125 IU hCG for luteal support), compared with the control group (hCG 6500 IU for trigger, vaginal P for luteal support).

FIG. 10

GnRH antagonist-based ovarian stimulation followed by GnRHa triggering and luteal support with 1500 IU of hCG when P level is below 30 nmol/L (luteal coasting).

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FIG. 11

Individual P changes following GnRHa in the early luteal phase. Based on data from Kol S, Breyzman T, Segal L, Humaidan P. ‘Luteal coasting’after GnRH agonist trigger— individualized, HCG-based, progesterone-free luteal support in ‘high responders’: a case series. Reprod Biomed Online 2015;31(6):747–751.

FIG. 12 GnRH antagonist-based ovarian stimulation followed by GnRHa triggering and luteal support with daily 125 IU of hCG from the day of oocyte retrieval until pregnancy test.

Modified Luteal Support With LH

Modified Luteal Support With GnRHa

Since recombinant LH has a shorter half-life compared with hCG, its use may achieve solid luteal support while potentially reducing OHSS risk. A small proof-of-concept study explored this strategy in “normal responders” [31]. Eighteen patients received GnRHa for trigger, followed by six doses of 300 IU recombinant LH (given on alternate days), and combined with standard support (Fig. 13). The control group received hCG for trigger (6500 IU) and standard support. Similar implantation rates were observed, with no OHSS cases in both groups. No further studies further explored this approach; therefore, its use in OHSS high-risk patients remains to be explored.

GnRHa trigger results in a deficient luteal phase, probably given its effect on LH secretion pattern as shown above [51]. Can repeated exposure to GnRHa during the luteal phase stimulate the corpora lutea? In a randomized prospective study, Pirard et al. [65] compared patients triggered with GnRHa (200 μg of nasal buserelin, study group), with patients triggered with 10,000 IU of hCG (control group). Luteal support in the study group was 100 μg nasal buserelin three times a day, and 200 mg vaginal P three times a day in the control group (Fig. 14). Luteal-phase P levels were comparable, except for higher levels on day 5 posttrigger day in the control

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FIG. 13 GnRH antagonist-based ovarian stimulation followed by GnRHa triggering and luteal support with six doses of 300 IU recombinant LH (given on alternate days from the day of oocyte retrieval) + standard LPS.

FIG. 14 GnRH antagonist-based ovarian stimulation followed by GnRHa triggering and luteal support with 100 μg nasal buserelin three times a day.

group. E2 levels were comparable during the entire luteal phase. Mean serum LH levels were significantly higher throughout the luteal phase in the study group than in the control group. Clinical outcome was similar. A similar approach was used by Bar-Hava et al. [66] in a retrospective cohort study of 46 high responders, ovulation was triggered with a single bolus of triptorelin (0.2 mg) and luteal support was initiated on the evening after oocyte retrieval with nafarelin inhaler (0.2 mg twice daily). High median P levels were measured at mid-luteal phase and on the day of the first positive pregnancy test (190 nmol/L in both measures). None of the patients developed OHSS, with an excellent clinical outcome (52.1% ongoing clinical pregnancies). This promising approach needs to be further explored.

Individualized Modified Luteal Support According to the Ovarian Response Any attempt to stimulate the corpora lutea following ovarian stimulation must be carefully considered in the context of potential OHSS. Individual response may vary considerably, even if all pertinent parameters (number and size of developing follicles, hormonal levels on trigger day, number of oocytes retrieved) are identical. Investigators have introduced guidelines and criteria to help clinicians reach a decision based on the number of developing follicles [57], or the combination of a few parameters [67]; however, the full scope of OHSS risk assessment is not limited to these factors. Therefore, caution and individual clinical judgment must be used to minimize OHSS incidence.

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GnRHa Trigger and “Freeze All” Since a nonsupplemented luteal phase post-GnRHa trigger will lead to complete luteal demise (in most cases), with nearly complete OHSS prevention, clinicians have introduced the option of “freeze-all,” or cycle segmentation. If a patient appears to hyperrespond, GnRHa trigger is administered, followed by oocyte retrieval and freezing of all embryos. In most cases, menses will begin within a week, after which a thaw cycle is performed, eliminating any OHSS risks. Significant improvements in cryobiology, especially vitrification, have questioned the advantage of a fresh vs thaw cycle. A Cochrane review [68] found no clear evidence of a difference in cumulative live birth rate between the freeze-all strategy and the conventional IVF/ICSI strategy. In a large (14,262 newborns) register-based cohort study [69], perinatal outcomes in children born after fresh or frozen ET were compared. In the autologous egg population, newborns from the fresh ET group had lower birthweight than the frozen-thawed ET group. In contrast, among egg-donor recipients undergoing ET, the mean birthweight did not differ between the groups. These findings suggest that fresh transfer might be involved with abnormal implantation secondary to endometrial exposure to excessive E2 levels during the follicular phase, and P levels during the early luteal phase. “Freeze all” may be dictated by the time frame imposed by preimplantation genetic screening. In this case, fresh transfer has no advantage over a frozen one as reported in a 179-patient randomized controlled study [70]. In fact, implantation rate per embryo transferred showed an improvement in the frozen group compared with the fresh group, but not significantly (75% vs 67%). Freezing all embryos allows for inclusion of all blastocysts in the cohort of embryos available for transfer, which also results in a higher proportion of patients reaching ET.

CONCLUSIONS GnRHa administration appears to be the preferable strategy for ovulation triggering in the context of OHSS prevention. Given its ability to induce LH and FSH surges, its use may be advantageous for all IVF cycles, replacing hCG as the gold standard trigger in IVF. If fresh transfer is sought in “high responders,” a wide range of strategies have been introduced to minimize the risk for OHSS; however, their use must be carefully considered against the full scope of individual clinical information. If it is anticipated that a fresh transfer puts the patient in significant risk to develop OHSS, “freeze-all” strategy is recommended. Clinical outcome in subsequent thaw cycles is not compromised.

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Glossary Corpus luteum (CL, or corpora lutea for plural) a robust ovarian endocrine tissue formed following oocyte extrusion from a mature follicle. CL produces progesterone that is crucial for a receptive endometrium. CL has a limited life span, unless rescued by hCG produced by the newly formed placenta. “Freeze all” freezing all embryos that developed after IVF treatment. Typically used in the context of OHSS prevention. GnRH antagonist-based ovarian stimulation protocols using GnRH antagonists during ovarian stimulation to prevent premature luteinization. LH (or FSH) surge a sharp rise in LH (or FSH) serum levels, that serves as a biochemical trigger of final oocyte maturation and ovulation. Luteal phase support hormonal treatment intended to support embryo implantation and pregnancy establishment. Luteolysis loss of corpus luteum hormonal function. Oocyte retrieval surgical intervention by which a needle is inserted into the ovarian tissue in order to aspirate follicles that contain oocytes. Ovarian hyperstimulation syndrome (OHSS) a medical syndrome that might develop following excessive ovarian stimulation. Mainly characterized by vascular fluid leakage through the pelvic capillary bed. Ovarian stimulation in IVF stimulating ovarian follicular growth with gonadotropins. Ovulation triggering medical induction of final oocyte maturation. “Segmentation” dissociate oocyte retrieval and embryo formation from embryo transfer. In most cases, embryo thaw and transfer is planned after a withdrawal bleeding that follows oocyte retrieval. Trophectoderm the outer layer of a blastocyst that will give rise to the placenta. Implantation window: a period (6–8 days postovulation or oocyte retrieval) during which an embryo can implant in the endometrium. Vitrification extremely rapid cooling of cell(s) or tissue to 196°C.

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[56] Griffin D, Benadiva C, Kummer N, Budinetz T, Nulsen J, Engmann L. Dual trigger of oocyte maturation with gonadotropin-releasing hormone agonist and low-dose human chorionic gonadotropin to optimize live birth rates in high responders. Fertil Steril 2012; 97(6):1316–20. [57] Humaidan P, Polyzos N, Alsbjerg B, Erb K, Mikkelsen A, Elbaek H, et al. GnRHa trigger and individualized luteal phase hCG support according to ovarian response to stimulation: two prospective randomized controlled multi-centre studies in IVF patients. Hum Reprod 2013;28:2511–21. det249. [58] Iliodromiti S, Blockeel C, Tremellen KP, Fleming R, Tournaye H, Humaidan P, et al. Consistent high clinical pregnancy rates and low ovarian hyperstimulation syndrome rates in high-risk patients after GnRH agonist triggering and modified luteal support: a retrospective multicentre study. Hum Reprod 2013;28:2529–36 det304. [59] Seyhan A, Ata B, Polat M, Son W-Y, Yarali H, Dahan MH. Severe early ovarian hyperstimulation syndrome following GnRH agonist trigger with the addition of 1500 IU hCG. Hum Reprod 2013;28:2522–8. det124. [60] O’Neill KE, Senapati S, Maina I, Gracia C, Dokras A. GnRH agonist with low-dose hCG (dual trigger) is associated with higher risk of severe ovarian hyperstimulation syndrome compared to GnRH agonist alone. J Assist Reprod Genet 2016;33 (9):1175–84. [61] Haas J, Kedem A, Machtinger R, Dar S, Hourvitz A, Yerushalmi G, et al. HCG (1500IU) administration on day 3 after oocytes retrieval, following GnRH-agonist trigger for final follicular maturation, results in high sufficient mid luteal progesterone levels-a proof of concept. J Ovarian Res 2014;7(1):35. [62] Delvigne A, Rozenberg S. Epidemiology and prevention of ovarian hyperstimulation syndrome (OHSS): a review. Hum Reprod Update 2002;8(6):559–77. [63] Kol S, Breyzman T, Segal L, Humaidan P. ‘Luteal coasting’ after GnRH agonist trigger—individualized, HCG-based, progesterone-free luteal support in ‘high responders’: a case series. Reprod Biomed Online 2015;31(6):747–51. [64] Andersen CY, Elbaek HO, Alsbjerg B, Laursen RJ, Povlsen BB, Thomsen L, et al. Daily low-dose hCG stimulation during the luteal phase combined with GnRHa triggered IVF cycles without exogenous progesterone: a proof of concept trial. Hum Reprod 2015; 30(10):2387–95. [65] Pirard C, Loumaye E, Laurent P, Wyns C. Contribution to more patient-friendly ART treatment: efficacy of continuous low-dose GnRH agonist as the only luteal support—results of a prospective, randomized, comparative study. Int J Endocrinol 2015; 2015:727569. [66] Bar-Hava I, Mizrachi Y, Karfunkel-Doron D, Omer Y, Sheena L, Carmon N, et al. Intranasal gonadotropin-releasing hormone agonist (GnRHa) for luteal-phase support following GnRHa triggering, a novel approach to avoid ovarian hyperstimulation syndrome in high responders. Fertil Steril 2016;106(2):330–3. [67] Papanikolaou EG, Humaidan P, Polyzos N, Kalantaridou S, Kol S, Benadiva C, et al. New algorithm for OHSS prevention. Reprod Biol Endocrinol 2011;9(1):147. [68] Wong KM, van Wely M, Van der Veen F, Repping S, Mastenbroek S. Fresh versus frozen embryo transfers for assisted reproduction. Cochrane Database Syst Rev 2017; Issue 3, Art. No.: CD011184. http://doi.org/10.1002/14651858.CD011184.pub2. [69] Vidal M, Vellve K, González-Comadran M, Robles A, Prat M, Torne M, et al. Perinatal outcomes in children born after fresh or frozen embryo transfer: a Catalan cohort study based on 14,262 newborns. Fertil Steril 2017;107(4):940–7. [70] Coates A, Kung A, Mounts E, Hesla J, Bankowski B, Barbieri E, et al. Optimal euploid embryo transfer strategy, fresh versus frozen, after preimplantation genetic screening with next generation sequencing: a randomized controlled trial. Fertil Steril 2017;107(3) 723–30.e3.

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C H A P T E R

24 The Ovarian Factor in Assisted Reproductive Technology Norbert Gleicher, Vitaly A. Kushnir, David H. Barad Abbreviations ACTH AFC AMH ART ASRM CDC COH DHEA DOR eSET ESHRE FOR FSH GC GnRH GSC hCG HGH hMG HPAA ICM IVF IU LFOR LH mDNA nDNA OHSS OI oPOI OR PB PCOS PGS PGT-A POA POR POI PVS SART SHBG SOI TSH

TOS ZP

adrenocorticotropic hormone antral follicle count anti-M€ ullerian hormone assisted reproductive technology American Society for Reproductive Medicine Center for Disease Control controlled ovarian hyperstimulation dehydroepiandrosterone diminished ovarian reserve elective single embryo transfer European Society for Human Reproduction and Embryology functional ovarian reserve follicle-stimulating hormone granulosa cell gonadotropin-releasing hormone germ-line stem cell human chorionic gonadotropin human growth hormone human menopausal gonadotropin hypothalamic-pituitary-adrenal axis inner cell mass in vitro fertilization international units low functional ovarian reserve luteinizing hormone mitochondrial DNA nuclear DNA ovarian hyperstimulation syndrome ovarian insufficiency occult primary ovarian insufficiency ovarian reserve polar body polycystic ovary syndrome preimplantation genetic screening preimplantation genetic testing for aneuploidy premature ovarian aging poor ovarian response primary ovarian insufficiency perivitilin space Society for Assisted Reproductive Technology sex hormone binding globulin secondary ovarian insufficiency thyroid-stimulating hormone

The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00024-8

total oocyte score zona pellucida

DEFINITIONS Definition of Ovarian Factor Embryos are typically created by successful fertilization of one oocyte by one spermatozoa. The resulting union of maternal and paternal nuclear haploid genomes (nDNA) establishes the newly diploid nDNA of the offspring embryo. It is always accompanied by an exclusively maternal mitochondrial genome (mDNA), as the paternal mitochondrial genome (spermatozoa) is lost soon after entering the maternal cytoplasmic oocyte microenvironment. Egg and sperm at fertilization, therefore, are not “equal” partners. As host-cell for the union of maternal and paternal genomes, the oocyte dominates the biology. Mitochondrial genetic diseases, consequently, are only inherited from mothers, and embryo quality is largely determined by oocytes rather than sperm. Among all contributing factors to reproductive success, the “ovarian factor” is, therefore, the most important and, likely, most multifaceted. It is defined by nDNA, mDNA, and other cytoplasmic constituents of importance; but also by folliculogenesis, the months-long maturation process of follicles and oocytes in the ovarian microenvironment, still by investigators not given appropriate attention. The “ovarian factor” is also highly age-dependent, still believed to be characterized by finite numbers of oocytes at their most primitive stage (in primordial follicles) in ovaries since embryonic life [some investigators have, although, raised the possibility of neoformation of follicles and oocytes from germ-line stem cells (GSCs)], and steadily declining in quantity and quality. As women age, oocytes are, therefore, presumed to “age” in parallel,

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reflected in decreasing functionality of nuclear and cytoplasmic components and in progressively declining egg quality. The presumed consequences are increases in meiotic chromosomal abnormalities, which increase with advancing female age from as low as 10%–20% to in excess of 60% [1]. We suggested a number of years ago a somewhat more complex concept of ovarian aging (the “CHR concept of ovarian aging”): primordial follicles at resting stage are primitive structures with almost no metabolic activity. Because they lack contact with their environment, they until recruitment, therefore, are in the external capsule of ovaries largely isolated from environmental influences. Only once recruited into folliculogenesis, do follicles/ oocytes establish increasing metabolic interdependence with their microenvironment and, therefore, become more vulnerable to environmental influences. This concept of ovarian aging, therefore, suggests that primordial follicles experience only minor damage while, often for decades, existing at resting stage in aging ovaries. Once recruited, however, they become subject to the toxic effect of an aging ovarian microenvironment, in which they undergo maturation over weeks to months. The damaging culprit in declining oocyte quality with advancing female age in this model is, therefore, not the time oocytes spent in the ovarian capsule as primordial follicles but the aging ovarian environment, in which they now must spend weeks to months of maturation, while migrating from the ovarian capsule inward toward the medulla [2]. Differences between above-described two hypotheses of oocyte aging are of considerable clinical importance: under the traditional hypothesis of ovarian aging, it is difficult to imagine that oocytes damaged by time may, still, be pharmacologically rescuable and repairable. Clinical interventions to improve oocyte quality in older women would, therefore, appear moot. Since annual US National Assisted Reproductive Technology (ART) outcome date demonstrate that reproductive endocrinologists rarely choose to treat women above age 42 with use of their own eggs, it is reasonable to assume that most of them still subscribe to this opinion. An aging ovarian microenvironment, however, could potentially still be subject to successful therapeutic interventions. The “CHR concept of ovarian aging” arose after observing beneficial effects of androgen supplementation on ovarian function, resulting in larger oocyte yield and improved oocyte quality [3,4]. This concept, therefore, recognizes the possibility that therapeutic interventions into early stages of folliculogenesis, which reconstitute physiological conditions of “younger” ovarian microenvironments and, therefore, improve the conditions of follicular maturation, may improve oocyte numbers and quality in older women. The “CHR concept of ovarian aging” creates the opportunity for pharmacological interventions into early stages of follicle maturation

to benefit older women who, still, wish to conceive with use of their own oocytes. Since males produce fresh sperm into advanced ages, paternal aging is of less concern, although recently reported data suggest that the decline in sperm quality with advancing male age may be more severe than has been so far appreciated.

Definition of ART ART encompassed a variety of treatments, techniques, and technologies. In this chapter, we will concentrate on the process of in vitro fertilization (IVF). IVF, normally, encompasses controlled ovarian hyperstimulation (COH) with fertility drugs, oocyte retrieval, IVF of oocytes with sperm in the laboratory, culture of resulting embryos, and embryo transfer into the uterus, either on day 3 after fertilization (cleavage stage) or on days 5–7 (blastocyst stage). COH in a large majority of cycles utilizes at different dosages gonadotropin stimulation, whether in form of injectable follicle-stimulating hormone (FSH) and/or human menopausal gonadotropin (hMG), which is a mixture of FSH and luteinizing hormone (LH). COH can also involve oral medications like clomiphene citrate and/or aromatase inhibitors, like letrozole, or a combination of orals and injectables. A small minority of IVF cycles utilize the natural cycle, thereby avoiding COH. Medication dosages are determined by a patient’s ovarian reserve (OR), which is generally understood as the estimated number of remaining follicles/oocytes in ovaries [2]. The lower the OR, the higher the required medication dosages that will be administered to obtain adequate ovarian responses to stimulation. Since OR in women declines with advancing age, medication requirements for COH usually increase as women age. A woman’s OR is made up of two distinct pools: a large majority of follicles/oocytes are in the so-called resting pool of primordial follicles, while at any given moment just a small proportion is in the so-called growing follicle pool (follicles after recruitment). Only the latter can be assessed with reasonable accuracy. In representation, and in contrast to the total OR that includes the resting follicle pool, we, therefore, describe this growing follicle pool in this chapter as the so-called functional ovarian reserve (FOR). Women with abnormally low numbers of growing follicles have low functional ovarian reserve (LFOR). The size of the resting follicle pool can be roughly estimated because it is usually proportional to the growing follicle pool [2]. Low OR (LOR) also called diminished OR (DOR) and LFOR are widely used terms, although, unfortunately, not well defined. It is important to recognize how important age is in determining what represents normal OR:

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what may be a perfectly normal OR at age 43, would be a LOR/LFOR at age 25. OR determinations should, therefore, be based on age-specific values of OR assessment tools, such as FSH, anti-M€ ullerian hormone (AMH), and antral follicle counts (AFCs) [2]. Another widely used term in the literature, mostly defined by the so-called Bologna criteria [5], is poor ovarian response (POR). Patients who produce smaller than expected oocyte yields are considered poor responders. Women with LOR/LFOR usually demonstrate POR. Since women become more resistant to ovarian stimulation with advancing age, like LOR/LFOR, POR also varies with female age. Although both represent distinctly different descriptions of ovarian function, LOR/LFOR and POR are often, nevertheless, used almost interchangeably. While diagnoses of LOR/LFOR and POR in most patients indeed overlap, they are clinically distinctively different: a diagnosis of LOR/LFOR (and, therefore, assumption of POR) can be reached in advance of even first IVF cycles; a diagnosis of POR, however, is only possible after at least one prior COH. This is, however, not the only reason why these two terms should be considered distinct. POR is an even more subjective diagnosis than LOR/LFOR since unexpectedly low responses to ovarian stimulation may also be caused by nonovarian causes. Specifically, iatrogenic interferences, like medication errors, wrong medication dosing and patient obesity may lead to a mistaken diagnosis of POR. LOR/LFOR, in contrast can always be assessed objectively by measuring FSH, AMH, and/or AFCs. Although the diagnosis of POR is widely used in the medical literature, we do not favor this diagnosis for either clinical or for study purposes, preferring the diagnosis of LOR/LFOR. Accurate and objective age-specific assessments of OR are essential in correctly defining the “ovarian factor” in infertile couples. Those then permit determinations whether age-specific ORs are normal, low, or high. Such determinations, in turn, often lead to diagnoses of premature ovarian aging (POA), also called occult primary ovarian insufficiency (oPOI) if OR is abnormally low or to the diagnosis of polycystic ovary syndrome (PCOS) if OR is unusually high [2]. Accurate OR assessments in advance are, of course, also of crucial importance in correctly individualizing and maximizing ovarian stimulation protocols for patients. The number of different ovarian stimulation protocols for IVF has proliferated in recent years, with some being of questionable efficacy [6]. A detailed review of all of them would exceed the framework of this chapter. Relevant details will, however, be discussed in —sections “What Controls the Ovary?,” “How the Ovary Controls Treatment Success in IVF,” and “Affecting Ovarian Performance.” Beyond new COH protocols, routine IVF over the last decade has undergone considerable changes. A variety of

so-called add-ons to standard IVF have been integrated, many increasingly questioned in their clinical utility [7]. Their discussion would also exceed the framework of this book chapter. Since so much depends on patient selection, outcome assessments in IVF are highly complex [8]. They, therefore, are often subject to subconscious or conscious manipulations [9]. When assessing IVF cycle outcomes, and one cannot review the “ovarian factor” in ART without paying close attention to IVF treatment outcomes, considerable care is, therefore, required to represent data correctly. Unfortunately, that has not always been the case in recent years, often misleading not only the public but also fertility practitioners. This chapter will, therefore, make special efforts to be transparent, and reflect IVF cycle outcomes correctly and objectively.

OVARIAN RESERVE Physiology of Ovarian Aging Fig. 1 demonstrates how a woman’s OR evolves from intrauterine life through menopause: peak follicle numbers in ovaries are reached at approximately 24 weeks intrauterine life, when ovaries contain approximately 7.0 million follicles. They then, however, rapidly decline in the steepest period of follicle loss to 3.5 million at time of birth, only to further diminish to 1300 female population controls, with odds ratio (OR) 1.30 per minor allele copy [26]. Subsequently, association of another FTO variant (rs1421085)

Ovarian Folliculogenesis Oligo-amenorrhea, anovulation, and impaired fertility are cardinal features of PCOS [33]. In a screen of 37 candidate genes, it was shown that 19p13.2 on chromosome 19 is strongly associated with susceptibility for development of PCOS [22,34]. Within 19p13.2, a dinucleotiderepeat polymorphism called D19S884 is known to map in close proximity to two genes of interest: intron 55 of FBN3 (the fibrillin 3 gene) and INSR (the insulin receptor gene). This polymorphism may influence the function of FBN3 (the third member of the fibrillin extracellular matrix protein family). It has been hypothesized that this in turn could impair follicle development in PCOS [22,34]. The function of INSR (and therefore insulin resistance) may also be influenced by this polymorphism. Gaining a clear understanding of genetic susceptibility to PCOS will provide much insight into its etiology and also provide a foundation on which to develop novel therapeutic strategies. The emergence of the GWAS has heralded a new era in the search for the genetic origins of PCOS. The GWAS is a much more powerful means of studying genetic susceptibility than the candidate gene approach, and one that has potential to identify

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EPIGENETIC MODIFICATIONS

unexpected and novel pathogenic pathways and therapeutic strategies. It is important that further GWAS in PCOS are conducted within larger and more diverse populations (including those of African ancestry), to gain further insight into the genetic and evolutionary origins of PCOS. However, alternative mechanisms of heritability should also be explored. Unfortunately, GWAS are blind to these alternative mechanisms of heritability. One such mechanism, and one which has attracted heightened interest in recent years, is that of “epigenetic modification.” It is this mechanism that we will explore in the next section of this chapter.

EPIGENETIC MODIFICATIONS The heritability of PCOS is analogous to matter in the universe. In this analogy, the baryonic matter that we can see (which accounts for 3400 subjects that compared with controls and following adjustment for differences in BMI, women with PCOS have lower levels of serum adiponectin [95]. It is known that adiponectin inhibits production of androgens from ovarian theca cells [96]. It follows therefore that suppressed levels of adiponectin may facilitate the effects of hyperinsulinemia on enhancement of ovarian androgen production [94]. Given the known effects of adiponectin on insulin sensitivity, lower adiponectin levels in PCOS probably also contribute toward insulin resistance that is characteristic in women with this condition [95]. In addition to effects on hyperandrogenism and reproductive function, weight gain induced insulin resistance in PCOS, as in metabolic syndrome [97], is also thought to be a major contributor to its associated cardiometabolic dysfunction. This includes the effects of insulin resistance on development of dyslipidemia [98], nonalcoholic fatty liver disease [4], and T2D [3]. Given the known association of PCOS with insulin resistance, obese women with PCOS often manifest heightened insulin resistance from the combined effects of PCOS and obesity: a metabolic “double-whammy” [2,6]. Such a scenario is likely to explain heightened risk of development of T2D (around 10%) and increased risk of early-onset impaired glucose tolerance (around 30%–40%) in women with PCOS [5]. Heightened insulin resistance in obese women with PCOS is also likely to underlie the increased risk of OSA in this condition: risk of OSA is 5- to 10-fold greater in women with PCOS compared with BMI-matched control women [5]. Our current understanding of the central role played by insulin resistance (and its worsening through weight gain) in the etiology of PCOS has an important implication for its effective management. Weight loss, through its beneficial effects on insulin resistance and therefore

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GUT MICROBIOTA

hyperinsulinemia, remains the most effective management strategy for obese women with PCOS. Benefits conferred from improved insulin sensitivity in PCOS include metabolic and ovulatory function, androgenicity, and fertility [2]. Lifestyle advice for obese women with PCOS has focused primarily on dietary modification [99]. Maintenance of weight loss remains a challenge for the longer-term management of PCOS [100]. This can be compounded in women with PCOS due to increased levels of low self-esteem, psychological distress, and depression that can interfere with the longer-term effective implementation of healthy lifestyle strategies [101]. Although bariatric surgery represents a promising solution, it is not a solution that is easily scalable to the population level. It is therefore important that research focus is maintained on development of novel, effective and durable weight-loss strategies in PCOS, including perhaps exploration of the effects of administration of novel weight-loss therapies. Our current therapeutic armamentarium for PCOS provides little more than symptomatic relief (with the exception of fertility treatments). We need to develop novel therapies for PCOS that address underlying etiological factors. Weight-related insulin resistance provides such a target.

GUT MICROBIOTA In recent years, gut microbiota have assumed increasing prominence with regard to their effects on general health, but particularly their relationship to chronic dysmetabolic conditions such as T2D. The human gut contains >100 trillion microbes, vastly outnumbering human cells [102]. Although we lack studies in women with PCOS, there are recent reports from rodent studies alluding to a potential role of gut microbiota in the development of PCOS. Many potential mutidirectional effects of gut microbiota may pertain (including involvement of the immune system, hormonal signaling [both levels of hormones and their receptor sensitivities], and nervous system [including effects on appetite and behavior]), but the “leaky gut hypothesis” has perhaps received most attention in recent years regarding effects on metabolic health. The leaky gut hypothesis purports that certain gut microbiota profiles (related to dietary and genetic factors) can predispose the colon to leakage of endotoxins (lipopolysaccharides produced by Gram-negative bacteria) from the gut into the serum [103]. Gut-derived endotoxinemia can then stimulate a chronic inflammatory response in the adipose and other tissues, which in turn can influence adipokine release and heighten insulin resistance and contribute to other metabolic disturbance [103]. Given the important role of insulin resistance in the etiology of PCOS [2,94], the potential role of gut

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microbiota in the development of this condition is an important consideration worthy of focused research. Guo and colleagues reported on the effects of gut microbiota in a rat model of PCOS [102]. Female Sprague-Dawley rats (n ¼ 32) were studied, with random assignment into four groups of eight rats each. These consisted of a control group and three PCOS groups (induced through letrozole treatment, an aromatase inhibitor that results in elevated levels of testosterone and suppressed estradiol levels). Fecal samples were collected, including at 21 days following letrozole treatment. One of the PCOS groups then underwent fecal microbiota transplantation (FMT) and another PCOS group underwent Lactobacillus transplantation. On day 36, fecal samples were collected and examined using real-time polymerase chain reaction (PCR). Real-time PCR and sequence analysis were used to analyze fecal microbiota. Hormonal levels of ostradiol and testosterone were also analyzed from blood samples and estrous cycle determination from vaginal smears. There were some important data generated. First, letrozole-induced PCOS was shown to influence gut microbiota: compared with fecal samples from control rats, those from letrozole-induced PCOS rats contained lower levels of Lactobacillus, Ruminococcus, and Clostridium and higher levels of Prevotella. In the PCOS rat groups administered FMT and Lactobacillus, the increased fecal Lactobacilli was associated with increases in estradiol. The associated reduction in fecal Prevotella in those fecal-transplanted PCOS rats was also associated with a reduction in serum testosterone and androstenedione. The fecal-transplanted rats also had improved estrous cycles and ovarian functions [102]. A further letrozole-induced PCOS rodent-based study on 4-week-old female mice reported by Kelley and colleagues also showed association of letrozole treatment with reduced microbiome diversity [104]. In this study, testosterone levels were inversely associated with diversity and abundance of large intestinal bacterial species. Letrozole treatment was associated with reduced fecal Bacteroidales and increased fecal Clostridiales and majority of Firmicutes [104]. As with the study reported by Guo and colleagues [102], the study outlined here also demonstrated an effect of letrozole-induced PCOS on reduced diversity of gut microbiota, presumed to be secondary to the hormonal changes (including elevated serum testosterone) induced by letrozole administration [104]. The studies outlined earlier show proof of concept that at least in a rodent model gut microbiota are implicated in reproductive function, and therefore may also play a role in the development of PCOS [102]. The interplay between gut microbiota and levels of sex hormones appears to be multidirectional. Lactobacillus is known to have health-promoting qualities and hypothesized to produce short-chain fatty acids within the colon that strengthen the gut barrier and reduce bacterial endotoxin

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translocation across the gut wall [105]. It has been shown that administration of Lactobacillus through probiotic supplements could reduce insulin resistance in humans in response to high-fat and calorie-laden diets [106]. More human-based studies on the role of fecal microbiota in the etiology of PCOS are required. Future study of fecal transplant or probiotic administration (including Lactobacillus) as a preventive and treatment strategy for women with PCOS should be prioritized, including effects of gut microbiota on insulin sensitivity and metabolic, biochemical, and clinical features of PCOS.

LACK OF EXERCISE AND PHYSICAL ACTIVITY Over recent decades, there has been mounting evidence reported in the literature to promote the health benefits of exercise and physical activity. Avoidance of adopting a sedentary lifestyle is topical and forms a cornerstone of general healthy living advice. It has been proposed that physical inactivity is an important cause of most chronic diseases [107]. Booth and colleagues argue, based on existing evidence from the literature, that physical activity prevents and delays onset of chronic disease affecting most systems that include metabolic, cardiovascular, musculoskeletal, gastrointestinal, psychological, and bone health [107]. Promotion of physical activity forms an important part of healthy lifestyle advice for women with PCOS. As such, exercise and physical activity are important environmental considerations when considering the pathogenesis of PCOS. Unfortunately, however, there is a lack of well-controlled studies reported in the literature that evaluate the benefits of exercise and physical activity both as a preventive and management strategy for PCOS [108]. The relevant literature is outlined here.

Reproductive Function There is some evidence that exercise can improve reproductive function in women with PCOS. In one study, restoration of normal menstrual cyclicity was shown in 60% of anovulatory women with PCOS who underwent a 3-month aerobic training program [112]. Menstrual cyclicity and ovulation rates also improved in overweight women with PCOS following 24 weeks of aerobic exercise [113]. In a further study that compared energy restricted diet alone or in combination with exercise, the combined group experienced more ovulatory cycles than the diet-alone group [114]. Overall, the data in the literature suggest that through exercise interventions, around 50% of women with PCOS will experience improvements in menstrual cyclicity and/or ovulation [108]. Although the mechanisms implicated are incompletely understood, it has been hypothesized that the reproductive benefits of exercise in PCOS are mediated, at least in part, through improvements in insulin resistance [108]. Although exercise appears to promote ovulation in women with PCOS, it should be noted that excessive exercise may have the opposite effect. In a systematic review of the literature on exercise and the effects on ovulation, Hakimi and colleague describe an increased risk of anovulation in extremely heavy exercisers (>60 min per day), but exercise duration 30–60 min per day appeared to reduce the risk of anovulatory infertility [115]. Excessive and heavy exercise (>60 min per day) may interfere with levels of leptin and opioids that in turn may affect both adrenal and gonadal function [115]. Promotion of moderate exercise and avoidance of excessive exercise may represent important health messages for women with PCOS.

Cardiovascular Health Physical Fitness A useful measure of physical fitness is the “VO2 max.” This is a measure of maximal aerobic capacity. There is controversy in the literature regarding VO2 max in women with PCOS. In one study, this measure of physical fitness is equivalent between women with PCOS and age- and BMI-matched controls [109], while another study showed an impairment of VO2 max in young overweight women with PCOS [110]. It has been proposed that such discrepancies of VO2 max in PCOS between studies could be influenced by differences in insulin sensitivity [108]. With regard to muscle strength [109] and levels of free-living physical activity [111], there appears to be no difference between women with PCOS and healthy controls.

In one study that included 90 overweight women with PCOS, a 3-month aerobic exercise program improved fasting insulin levels, BMI, and waist circumference compared with nonexercising controls [112]. In a comparison between 24 weeks of aerobic exercise vs a hypocaloric diet in 40 obese women with PCOS, the exercise group had greater improvements in waist circumference, insulin levels, and free androgen index [113]. In a more detailed and complex study that included comparisons of dietalone, with diet combined with different forms of exercise (aerobic and aerobic-resistance) over a 20-week period in obese and overweight women with PCOS, there were improvements in body weight, blood pressure, lipid profile, glucose, and insulin levels in all groups [114]. However, the combined exercise groups experienced more favorable changes in body composition than the diet-alone

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group, with relative reductions in fat mass and preservation of fat-free mass [114].

Psychological Well-Being There are few studies that report psychological effects of physical activity in PCOS. Body image distress score was improved in women with PCOS following a briskwalking program that was self-directed for a duration of 6 months [116]. In a broader cohort of obese infertile women who followed lifestyle modification programs (that included increased exercise), there were improvements in self-esteem, depression, and anxiety [117]. More recently, Banting and colleagues published self-reported measures of depression and anxiety in women with and without PCOS. Regardless of PCOS status, physically active women had less severe depression than their inactive counterparts [118]. Such associative data are difficult to interpret given that causality cannot be inferred, and multidirectional effects could pertain. Although women with PCOS identified more sources of support than the control women, there were more barriers to exercise for women with PCOS compared with controls (including lack of confidence, fear of injury, and physical limitations) [118]. To summarize this section, although the literature is limited in this field, there does appear to be good evidence to support the reproductive (including menstrual cyclicity and ovulatory rate) and cardiovascular (including insulin resistance and body composition) health benefits conferred by moderate exercise and physical activity in overweight and obese women with PCOS. There are likely to be many other benefits of exercise in PCOS (as in the general population) that include psychological well-being, although direct evidence for this is limited due to paucity of published relevant studies. Physical fitness and muscle strength per se do not appear to be impaired in PCOS, although in some women with this condition, other impediments to exercise may pertain (such as reluctance to attend a gym due to impaired selfesteem, for example). An important learning point from the available literature is that exercise and physical activity in PCOS (as in the general population) appear to confer health benefits that are independent of dietary change and that these benefits occur even without weight loss. This important public health message needs to be promulgated widely: although weight loss is often an important target in obese women with PCOS, significant improvements in reproductive and cardiovascular health can also be achieved simply through increased exercise and physical activity, even without attendant weight loss. Optimal lifestyle change of course combines dietary change with increased exercise and physical activity, which in most cases would also be expected to result in some weight loss.

Finally, much of the discussion in this section has been focused on studies on exercise and physical activity in women with an established diagnosis of PCOS: severity of reproductive and cardiovascular manifestations of PCOS appears to be influenced by physical activity. The role of physical activity (or lack thereof) as an “environmental” (i.e., nongenetic) contributor to future development of PCOS has not been demonstrated conclusively in any longer-term prospective study. However, given the well-established contribution of weight gain in the development of PCOS in women who are genetically predisposed [2], and the known contribution of lack of exercise on weight gain and insulin resistance [119], it seems reasonable to hypothesize that female sedentariness, especially when associated with weight gain, would likely be an important environmental contributor to future development of PCOS.

SOCIOECONOMIC STATUS It is well-established that lower socioeconomic status (SES) associates with engagement in unhealthy lifestyles that include poor nutrition, lack of physical activity, and smoking [120]. There is also an association between low SES and obesity in women [121]. Given the link between obesity and PCOS [2], it is important to consider the role of SES as an environmental contributor to the etiology of PCOS. Merkin and colleagues studied women (n ¼ 1163) from the Coronary Artery Risk Development in Young Adults (CARDIA) Women’s Study, a prospective study of risk factors for coronary artery disease in young adults [122]. Self-reported data on menstrual cyclicity and hirsutism were ascertained, and testosterone and SHBG measured. Parental education was used as a marker of childhood SES, and personal education of each respondent as a measure of adult SES. SES trajectories were constructed for each subject, based on childhood and adult SES scores (4 trajectories with SES for each marked as “high” or “low”). Within the cohort, the overall prevalence of PCOS was 10.7%. Those women with lower childhood SES scores tended to have a higher prevalence of PCOS development. Interestingly, the subgroup with both low childhood SES and high adult SES appeared to be most at risk of developing PCOS [122]. It was hypothesized by the authors that compared with those with a low adult SES, women with high adult SES may have better recall of features relating to menstruation and hirsutism, and therefore may be more likely to have a diagnosis of PCOS. Consistent with this hypothesis is the observation that serum testosterone had the strongest correlation with PCOS in the subgroup of women with both low childhood and adult SES [122]. Another proposed possible explanation for the data is that upwardly

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mobile women may experience increased peripubertal stress that in turn could contribute toward weight gain and menstrual irregularity [122]. Although self-reporting was a limitation of the study reported by Merkin and colleagues [122], a clear strength was the size of the cohort and the availability of biochemical data. The effects of SES mobility and transition in the peripubertal period on metabolic health and future risk of PCOS development should be a focus for future research. It is fairly clear from the study reported [122], however, that low childhood SES is associated with increased risk for future development of PCOS. It has been suggested that poor intrauterine or childhood nutrition may drive association between low childhood SES and risk of developing PCOS [123]. Other possible explanations for this association include childhood adoption of poor healthrelated behaviors that persist into adulthood [122] and associated excessive weight gain in infancy and early childhood [124]. Intriguingly, the effects of SES on risk for development of PCOS may be dependent upon the population studied and its prevailing cultural and economic milieu. While studies in western countries reveal an association between low SES in childhood and future development of PCOS, as exemplified by Merkin and colleagues [122], the situation in those countries that over recent decades have undergone, and are undergoing rapid westernization may be different. In such regions of the world such as much of the Indian subcontinent, affluence rather than poverty is likely to be an important driver of weight gain and associated metabolic dysfunction. In one study on women from Mumbai in India, PCOS developed in predominantly middle-class populations: presumably those who could afford to adopt a “western-type” lifestyle with the unfortunate associated weight gain [125]. The observation by Pathak and colleagues [125], and the juxtaposition with PCOS prevalence and SES status in western countries, reveals a profound insight: ultimately, caloric intake is likely to mediate association between SES and PCOS development. In poorer populations, affluence may provide a key to excessive caloric ingestion which the poorer members of those societies can simply not afford. In more affluent populations, relative poverty may drive excessive caloric ingestion through relative affordability of cheaper and unhealthy “convenience” foods. Although there are likely many other lifestyle factors at play, the effects of SES on caloric ingestion and PCOS development in different populations are important considerations. In addition to risk for development of PCOS, SES may also influence the phenotypic expression of PCOS. In one study of 244 women with PCOS who completed a questionnaire regarding family income and school education, Di Fede and colleagues reported an association between income and education with ovulation (assessed through

measurement of day 22 serum progesterone following a spontaneous or induced menstrual cycle) [126]. The proportion of women with PCOS who manifested ovulation was lower in those with low-medium income (21%) vs those with high income (43%). Similarly, those women with PCOS who had low education also had lower rates of ovulation (12%) compared with the subgroup with high education level (47%) [126]. The authors hypothesized that the differences in ovulation between the subgroups are likely related to differences in BMI and insulin levels (with an inverse correlation between family income and BMI and insulin levels and an inverse relationship between insulin and progesterone levels) [126]. SES as an entity is difficult to study based on a number of factors. These include differing definitions between studies on different populations in different cultures, and also the transitory nature of SES, particularly in our modern-day environment, with upward trajectories fairly commonplace particularly in rapidly westernized populations. A further difficulty with SES however, other than its definition and transitory nature, is that SES is often intertwined with, and has complex interactions with a person’s lifestyle and behavior, for a multitude of reasons. Any association with SES is therefore likely to be complex and manifest multidirectionality. Therefore, any discussion of epidemiological data regarding SES and PCOS is necessarily associative by nature, and firm conclusions regarding SES as a causal environmental factor in the etiology of PCOS are inevitably limited. Notwithstanding these limitations, however, the available data reveal population-specific associations of SES with development of PCOS. While low SES in childhood appears to associate with later development of PCOS, in poorer populations, the reverse may pertain, with higher SES and associated affluence associated with weight gain and increased risk for development of PCOS. As outlined, phenotypic subgroup of PCOS may also be influenced by SES. It is important to focus on the actual mechanisms that mediate effects of SES on PCOS development in each specific population and translate these into meaningful and targeted public health messages. Instead of focusing primarily on SES per se, focus should instead rest on targeting those SES groups most at risk in any given population, and facilitating change toward a more salutary lifestyle for all. Focused and SES-relevant dietary advice should form an important part of this initiative.

ADVANCED GLYCATION END PRODUCTS In recent years, there has been much interest in the deleterious metabolic effects of a diverse group of reactive molecules called advanced glycation end products (AGEs), through a plethora of possible mechanisms that

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ADVANCED GLYCATION END PRODUCTS

includes activation of a chronic inflammatory response and cellular damage [127]. AGEs are formed endogenously through nonenzymatic advanced glycation and oxidation (glycoxidation) reactions of lipids, nucleic acids, and free amino groups of proteins with carbohydrates [128]. AGEs also occur in our food, particularly cooked “fast-food” diets, uncooked animal-derived foods, high protein and fat content, and thermal processing of foods [129]. Dietary intake, together with tobacco represents an important source of exogenous AGEs. Evidence has emerged in recent years to implicate a potential role for dietary AGEs as an environmental contributor to the etiology of PCOS, including its dysmetabolic and reproductive features [128,130]. It is known that the harmful proinflammatory effects of AGE are mediated via their interaction with the receptor for advanced glycation end products (RAGE), which are transmembrane receptors. AGE-RAGE interaction results in a proinflammatory response, oxidative stress (including activation of nuclear kappa B [NF-κB]), and ultimately tissue damage [128]. AGE-RAGE interaction also upregulates RAGE expression acting like a positive feedback loop [128,131]. Conversely, presence of soluble receptor for advanced glycation end products (sRAGE) acts like a brake on this mechanism and confers protection from the harmful effects of AGE through antiinflammatory effects [128]. Unlike RAGE, sRAGE circulates throughout the body and forms a decoy receptor that can bind to circulating AGEs, thereby preventing the harmful effects of AGE-RAGE interaction [128]. It has been reported that circulating levels of AGE and ovarian RAGE are elevated in women with PCOS [128,132]. Furthermore, ingestion of a diet that is low in AGE-content in women with PCOS has been shown to be associated with improved hormonal and metabolic profiles [128]. It has been demonstrated that serum levels of AGE (from dietary intake) actually correlate with inflammatory markers that include C-reactive protein and HOMA IR (a marker of insulin resistance) [128]. Production of reactive oxygen species within tissues following accumulation of dietary AGEs can enhance cellular damage [133]. PCOS is a condition characterized by chronic low-grade inflammation, which likely underlies some of the metabolic and reproductive features of this condition [134]. It has been hypothesized that dietary AGEs may contribute to the chronic inflammatory state of PCOS, and may therefore contribute toward its pathogenesis [128]. However, further studies are required to validate this hypothesis. One type of AGE is termed methylglyoxal (MG). MG is metabolized by an enzyme called Glyoxalase 1 (GLO-1), thereby providing some protection against the deleterious effects of AGEs [135]. It has been demonstrated in Wistar rats that ovarian accumulation of AGE results from reduced ovarian activity of GLO-1, which in turn

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can result from increased dietary intake of AGEs combined with excessive exposure to androgens within the ovary [136]. It has been hypothesized that such a mechanism may result in elevated inflammatory markers within the ovary, which in turn may contribute toward ovarian dysfunction that typifies PCOS [136]. There is accumulating evidence from the literature to support deleterious effects of AGE-RAGE on folliculogenesis, oocyte maturation [137], mitochondrial function, apoptosis, and DNA damage within the oocyte [137]. There is evidence to suggest that a diet low in AGE content in women with PCOS (compared with high AGE content) is associated with beneficial effects on metabolic factors and oxidative stresses [138]. In one study, women with PCOS were fed a high-AGE vs a low-AGE diet for 2 months, with marked improvement in insulin sensitivity (HOMA IR marker) following the low-AGE diet [138]. It has been demonstrated in vitamin D-deficient women with PCOS that dietary vitamin D supplementation results in increased serum sRAGE [139]. Furthermore, metformin intake for 6 months in women with PCOS has been shown to result in a reduction in serum AGE levels [140]. It is believed that metformin reduces expression of RAGE, thereby preventing AGE-induced cell damage [141]. Metformin may also convert MG to dihydroimidazole, thereby reducing oxidative stress [140]. To summarize, there is evidence in the literature from a variety of sources that include animal models of PCOS, in vitro studies and studies on women with PCOS, to support association between dietary AGE intake and deleterious inflammatory and metabolic effects (including cellular damage from oxidative stress) and ovulatory dysfunction. However, much of the literature in this field is based on associative data, from which we should be cautious about inference of causality. Although a plausible causal mechanism between AGE-RAGE and inflammatory, metabolic, and reproductive features of PCOS has been hypothesized, further focused studies are required to confirm or refute such a causal mechanism, to elucidate which, if any pathways link precisely AGERAGE with PCOS pathogenesis, and how such insight can be used to develop potential novel therapies. Given the association of high AGE content in food with unhealthy “fast food,” it remains difficult to disentangle the harmful metabolic effects of fat and carbohydrate in food highly laden with these macronutrients, from those of AGE content per se. However, despite our incomplete understanding of causality, on the basis of current evidence, it would seem a reasonable approach for general healthy lifestyle advice to include limitation of our intake of dietary AGEs and advice on healthier cooking methods: AGE content of food is limited through preparation of food at low temperature, with high moisture, brief heating time, and use of acidic marinades (including lemon juice and vinegar) [128].

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Low maternal socio economic status

Fetal development (fetal programming hypothesis)

Maternal under-and poor-nutrition

Fetal nutritional insufficiency

Endocrine-disrupting chemicals Maternal-fetal exposure to: Triclocarbon Bisphenol A Phthalates Perfluoroalkyl acids Nicotine

Fetal excess androgen exposure

Epigenetic effects Fetal insulin resistance Tissue programming

Cardinal features of PCOS Excessive ovarian theca cell androgen production

Impaired follicular development

Gut microbiota and advanced glycation end products

Vitamin D deficiency Inadequate UVB exposure Poor diet

Chronic inflammatory response

Reduced serum sRAGE levels

Insulin resistance and secondary hyperinsulinaemia

Gut-derived endotoxinaemia Prevotella Lactobacillus Poor diet Cooking methods AGE/RAGE complexes

Sequelae of obesity

Population-specific adult socioeconomic status

Genetic and epigenetic factors Weight gain

FTO variants

Poor diet Caloric excess Lack of physical activity

Poor well-being

FIG. 2 Summary of the mechanisms by which environmental factors are implicated in the etiology of PCOS (where relevant, genetic, and epigenetic factors are also shown).

A summary of mechanisms by which environmental factors are implicated in the etiology of PCOS is shown in Fig. 2.

CONCLUSIONS As the burgeoning obesity epidemic ensues, the prevalence of PCOS around the world is likely to increase. PCOS is a condition associated with significant physical and mental morbidity, and one which, potentially, confers a substantial cardiovascular burden. A fundamental premise of any human condition is to understand its etiology. Unfortunately, despite decades of focused research across many fields, our understanding of the

etiology of PCOS remains incomplete. There are many possible reasons for this, which include the complex and heterogeneous nature of PCOS and the inherent difficulties associated with its effective study as outlined earlier. However, despite these barriers, we must not let these detract us from striving in the future to gain a clear understanding of PCOS etiology. In this chapter, we have explored the three main components of the etiology of PCOS: genetic, epigenetic, and environmental. Within each section, the relevant published data have been discussed, including implications for effective management, and future directions for further research. It is of course possible that other hitherto unknown components of PCOS etiology play contributory roles and these will become evident as data from

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further studies emerge. Although we understand heritability as a statistical likelihood of developing PCOS, and the roles of weight gain and insulin resistance in the expression and manifestation of PCOS, we still lack a clear and detailed understanding of the underlying mechanisms at play. A general observation from the PCOS literature is that there seems to be a predominance of cross-sectional studies reported. Unfortunately, such studies are inherently limited in their capacity to shed insight into underlying mechanisms, providing only a “snap-shot” vista. To truly understand mechanisms, we need more long-term prospective studies that explore the factors (genetic, epigenetic, and environmental) that may influence the clinical course of PCOS in women with this condition over time. Such studies will provide important insights into all the etiological pathways discussed in this chapter. We may never fully understand the true complexity of PCOS and its intricate etiology. However, improved understanding of the etiology of PCOS will enable us to manage this condition more effectively. Our ability to accurately diagnose, predict, and prevent the onset of PCOS is also likely to improve. Such measures will hopefully reduce both the prevalence and the burden and morbidity on the many women and girls, globally, who suffer from PCOS.

Acknowledgments We acknowledge the many patients, relatives, nurses, and physicians who contributed to the ascertainment of the various clinical samples reported on in this chapter.

Conflicts of Interest None.

Financial Disclosure None.

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[58] Yin M, Wang X, Yao G, Lu M, Liang M, Sun Y, et al. Transactivation of micrornA-320 by microRNA-383 regulates granulosa cell functions by targeting E2F1 and SF-1 proteins. J Biol Chem 2014; 289(26):18239–57. [59] Chen YH, Heneidi S, Lee JM, Layman LC, Stepp DW, Gamboa GM, et al. miRNA-93 inhibits GLUT4 and is overexpressed in adipose tissue of polycystic ovary syndrome patients and women with insulin resistance. Diabetes 2013;62(7):2278–86. [60] Sir-Petermann T, Hitchsfeld C, Maliqueo M, Codner E, Echiburu B, Gazitua R, et al. Birth weight in offspring of mothers with polycystic ovarian syndrome. Hum Reprod 2005;20 (8):2122–6. [61] Dumesic DA, Schramm RD, Abbott DH. Early origins of polycystic ovary syndrome. Reprod Fertil Dev 2005;17(3):349–60. [62] Gur EB, Karadeniz M, Turan GA. Fetal programming of polycystic ovary syndrome. World J Diabetes 2015;6(7):936–42. [63] Abbott DH, Barnett DK, Bruns CM, Dumesic DA. Androgen excess fetal programming of female reproduction: a developmental aetiology for polycystic ovary syndrome? Hum Reprod Update 2005; 11(4):357–74. [64] Dumesic DA, Schramm RD, Bird IM, Peterson E, Paprocki AM, Zhou R, et al. Reduced intrafollicular androstenedione and estradiol levels in early-treated prenatally androgenized female rhesus monkeys receiving follicle-stimulating hormone therapy for in vitro fertilization. Biol Reprod 2003;69(4):1213–9. [65] Kandaraki E, Chatzigeorgiou A, Livadas S, Palioura E, Economou F, Koutsilieris M, et al. Endocrine disruptors and polycystic ovary syndrome (PCOS): elevated serum levels of bisphenol A in women with PCOS. J Clin Endocrinol Metab 2011;96(3): E480–4. [66] Dumesic DA, Abbott DH, Padmanabhan V. Polycystic ovary syndrome and its developmental origins. Rev Endocr Metab Disord 2007;8(2):127–41. [67] Abbott DH, Dumesic DA, Franks S. Developmental origin of polycystic ovary syndrome—a hypothesis. J Endocrinol 2002;174(1): 1–5. [68] Hewlett M, Chow E, Aschengrau A, Mahalingaiah S. Prenatal exposure to endocrine disruptors: a developmental etiology for polycystic ovary syndrome. Reprod Sci 2017;24(1):19–27. [69] Halden RU. On the need and speed of regulating triclosan and triclocarban in the United States. Environ Sci Technol 2014;48(7): 3603–11. [70] Chen J, Ahn KC, Gee NA, Ahmed MI, Duleba AJ, Zhao L, et al. Triclocarban enhances testosterone action: a new type of endocrine disruptor? Endocrinology 2008;149(3):1173–9. [71] Pycke BF, Geer LA, Dalloul M, Abulafia O, Jenck AM, Halden RU. Human fetal exposure to triclosan and triclocarban in an urban population from Brooklyn, New York. Environ Sci Technol 2014;48(15):8831–8. [72] Ikezuki Y, Tsutsumi O, Takai Y, Kamei Y, Taketani Y. Determination of bisphenol A concentrations in human biological fluids reveals significant early prenatal exposure. Hum Reprod 2002; 17(11):2839–41. [73] Heudorf U, Mersch-Sundermann V, Angerer J. Phthalates: toxicology and exposure. Int J Hyg Environ Health 2007;210 (5):623–34. [74] Wittassek M, Angerer J, Kolossa-Gehring M, Schafer SD, Klockenbusch W, Dobler L, et al. Fetal exposure to phthalates— a pilot study. Int J Hyg Environ Health 2009;212(5):492–8. [75] Pocar P, Fiandanese N, Secchi C, Berrini A, Fischer B, Schmidt JS, et al. Exposure to di(2-ethyl-hexyl) phthalate (DEHP) in utero and during lactation causes long-term pituitary-gonadal axis disruption in male and female mouse offspring. Endocrinology 2012;153(2):937–48.

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[113] Palomba S, Giallauria F, Falbo A, Russo T, Oppedisano R, Tolino A, et al. Structured exercise training programme versus hypocaloric hyperproteic diet in obese polycystic ovary syndrome patients with anovulatory infertility: a 24-week pilot study. Hum Reprod 2008;23(3):642–50. [114] Thomson RL, Buckley JD, Noakes M, Clifton PM, Norman RJ, Brinkworth GD. The effect of a hypocaloric diet with and without exercise training on body composition, cardiometabolic risk profile, and reproductive function in overweight and obese women with polycystic ovary syndrome. J Clin Endocrinol Metab 2008;93(9):3373–80. [115] Hakimi O, Cameron LC. Effect of exercise on ovulation: a systematic review. Sports Med 2017;47:1555–67. [116] Liao LM, Nesic J, Chadwick PM, Brooke-Wavell K, Prelevic GM. Exercise and body image distress in overweight and obese women with polycystic ovary syndrome: a pilot investigation. Gynecol Endocrinol 2008;24(10):555–61. [117] Clark AM, Thornley B, Tomlinson L, Galletley C, Norman RJ. Weight loss in obese infertile women results in improvement in reproductive outcome for all forms of fertility treatment. Hum Reprod 1998;13(6):1502–5. [118] Banting LK, Gibson-Helm M, Polman R, Teede HJ, Stepto NK. Physical activity and mental health in women with polycystic ovary syndrome. BMC Womens Health 2014;14(1):51. [119] Chaput JP, Tremblay A. Obesity and physical inactivity: the relevance of reconsidering the notion of sedentariness. Obes Facts 2009;2(4):249–54. [120] Barkley GS. Factors influencing health behaviors in the National Health and Nutritional Examination Survey, III (NHANES III). Soc Work Health Care 2008;46(4):57–79. [121] Thurston RC, Kubzansky LD, Kawachi I, Berkman LF. Is the association between socioeconomic position and coronary heart disease stronger in women than in men? Am J Epidemiol 2005; 162(1):57–65. [122] Merkin SS, Azziz R, Seeman T, Calderon-Margalit R, Daviglus M, Kiefe C, et al. Socioeconomic status and polycystic ovary syndrome. J Womens Health (Larchmt) 2011;20(3):413–9. [123] Davey Smith G, Hart C. Insulin resistance syndrome and childhood social conditions. Lancet 1997;349(9047):284–5. [124] Gardner DS, Hosking J, Metcalf BS, Jeffery AN, Voss LD, Wilkin TJ. Contribution of early weight gain to childhood overweight and metabolic health: a longitudinal study (EarlyBird 36). Pediatrics 2009;123(1):e67–73. [125] Pathak G, Nichter M. Polycystic ovary syndrome in globalizing India: an ecosocial perspective on an emerging lifestyle disease. Soc Sci Med 2015;146:21–8. [126] Di Fede G, Mansueto P, Longo RA, Rini G, Carmina E. Influence of sociocultural factors on the ovulatory status of polycystic ovary syndrome. Fertil Steril 2009;91(5):1853–6. [127] Ulrich P, Cerami A. Protein glycation, diabetes, and aging. Recent Prog Horm Res 2001;56:1–21. [128] Garg D, Merhi Z. Advanced glycation end products: link between diet and ovulatory dysfunction in PCOS? Nutrients 2015;7(12): 10129–44. [129] Goldberg T, Cai W, Peppa M, Dardaine V, Baliga BS, Uribarri J, et al. Advanced glycoxidation end products in commonly consumed foods. J Am Diet Assoc 2004;104(8):1287–91. [130] Diamanti-Kandarakis E, Dunaif A. Insulin resistance and the polycystic ovary syndrome revisited: an update on mechanisms and implications. Endocr Rev 2012;33(6):981–1030. [131] Diamanti-Kandarakis E, Katsikis I, Piperi C, Kandaraki E, Piouka A, Papavassiliou AG, et al. Increased serum advanced glycation endproducts is a distinct finding in lean women with polycystic ovary syndrome (PCOS). Clin Endocrinol (Oxf) 2008;69(4):634–41. [132] Diamanti-Kandarakis E, Piperi C, Patsouris E, Korkolopoulou P, Panidis D, Pawelczyk L, et al. Immunohistochemical localization of advanced glycation end-products (AGEs) and their receptor

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C H A P T E R

28 Germ Cell Failure and Ovarian Resistance: Human Genes and Disorders Joe Leigh Simpson, Yingying Qin, Zi-Jiang Chen INTRODUCTION Ovarian failure and ovarian resistance in humans have a variety of causes. Failure may be complete or premature, occurring earlier (younger than 35 or 40 years) than the expected age of menopause. Either complete or premature ovarian failure (POF) has long been deduced to result from either deletions of the X chromosome or Mendelian causes; however, only rarely is the actual gene known. In this discussion, we shall systematically survey the known disorders of ovarian failure and ovarian resistance in humans. This chapter inevitably reflects our previous publications.[1]

EMBRYOLOGY OF OVARIAN DEVELOPMENT: NATURE OF GENES TO BE POSTULATED Primordial germ cells (PGCs) originate in the endoderm of the yolk sac and migrate to the genital ridge to form the indifferent gonad. 46,XY and 46,XX gonads are initially indistinguishable. Indifferent gonads develop into testes if the embryo, or more specifically the gonadal stroma, is 46,XY. Testes become morphologically identifiable 7–8 weeks following conception (9–10 week gestational or menstrual weeks). Ovaries become identifiable thereafter. In the absence of a Y chromosome, the indifferent gonad develops into an ovary. Transformation into fetal ovaries begins at 50 and 55 days of embryonic development. It is debatable whether female (ovarian) differentiation is truly a default (constitutive) pathway, or whether a specific gene product directs primary ovarian differentiation. At one time the default hypothesis

The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00028-5

seemed most plausible, but more recently a primary directive role in ovarian differentiation has been proposed anew. Previously, an attractive candidate gene for this role was DAX-1 (dosage-sensitive sex reversal/adrenal hypoplasia critical region X). DAX-1 was attractive because it was shown to be encoded in the region of the X short arm (Xp21), which could redirect 46,XY embryos into female differentiation when duplicated. The mouse homolog for human AHC was Ahch. If Ahch (DAX1) were to play a pivotal role in primary ovarian differentiation, Ahch should be upregulated in the XX mouse ovary. This did occur, and transgenic XY mice overexpressing Ahch developed as females, at least in the presence of a relatively weak Sry; however, XX mice lacking Ahch unexpectedly showed ovarian differentiation, ovulated and were even fertile! [2] Thus Ahch cannot be responsible for primary ovarian differentiation in mice, nor presumably is DAX1 in (AHC) humans. Irrespective of the aforementioned uncertainty, the key observation—clinically and genetically—is that germ cells are present in 45,X embryos. This can be deduced by the presence of germ cells in 45,X abortuses, which account for 10% of all first-trimester spontaneous abortions. The pathogenesis of germ-cell absence in 45,X adults thus involves atresia, occurring at a rate more rapid than that occurring in normal 46,XX embryos. Pathogenesis does not involve failure of germ-cell formation. In fact, germ cells are present in virtually all monosomy X mammals (e.g., mice) and often adults as well. If two intact X chromosomes are required to prevent human 45,X ovarian follicles from degenerating prematurely, it follows that the second X chromosome in humans must be responsible for ovarian maintenance, rather than primary ovarian differentiation.

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OVARIAN FAILURE AS A RESULT OF HAPLOINSUFFICIENCY FOR THE X CHROMOSOME (X-MONOSOMY AND X-DELETIONS)

TABLE

1 Somatic Features Associated Chromosomal Complement

With

the

45,X

Growth Decreased birth weight Decreased adult height (141–146 cm)

Nomenclature

Intellectual function

The first group of subjects with ovarian failure was recognized in 1938 by Turner. These women had not only ovarian failure but also short stature and other now wellknown somatic anomalies. For many years, the term Turner syndrome was applied to all women with ovarian failure. Yet Turner syndrome is a broad term, and we prefer to apply the term gonadal dysgenesis to women with streak gonads and reserve the term Turner stigmata for those having short stature and certain somatic anomalies (Table 1). By itself, Turner stigmata as defined would not imply the presence of streak gonads. The term Turner syndrome would be applied to those individuals with streak gonads, Turner stigmata, and a 45,X or X-deletion complement.

Verbal IQ > performance IQ Cognitive deficits (space-form blindness) Immature personality, probably secondary to short stature Cranofacial Premature fusion sphenoccipital and other sutures, producing brachycephaly Abnormal pinnae Retruded mandible Ptosis hypertelorism Epicanthal folds (25%) High-arched palate (36%) abnormal dentition Visual anomalies, usually strabismus (22%), amblyopia hypernetropia auditory deficits; sensorineural or secondary To middle ear infection Neck Pterygium colli (46%) short, broad neck (74%) low nuchal hair (71%)

Monosomy X

Chest

Monosomy X explains perhaps 50% of hypergonadotropic hypogonadism. In 80% of 45,X cases, the X is maternal (Xm) in origin. In the remaining 20%, the remaining X is paternal (Xp) in origin. That 45,X individuals show streak gonads as adults is not as obvious as might be expected because relatively normal ovarian development occurs in many other mammals (e.g., mice) with monosomy X. The presumptive explanation is that pivotal genes on the heterochromatic (inactive) X are not inactivated. Of the some 2000 genes on the X, perhaps 5% escape inactivation. Most of these genes are on the X short arm (Xp), clustered in selected euchromatic regions. Candidate genes for ovarian maintenance genes probably lie in these euchromatic regions although studies have not truly addressed this obvious postulate. Not all 45,X patients show complete ovarian failure. Approximately 3%–5% of adult 45,X patients menstruate spontaneously. Fertile patients have been reported, as reviewed by Abir et al. [3] and Hovatta [4]. An undetected 46,XX cell line (i.e., 45,X/46,XX mosaicism) should be suspected in menstruating 45,X patients, which seems especially plausible in some reports. Magee et al. [5] observed seven pregnancies in one ostensibly 45,X woman; however, it is not unexpected that some 45,X individuals could be fertile, inasmuch as germ cells are present in 45,X embryos.

Rectangular contour (shield chest) (35%) apparent widely spaced nipples Tapered lateral ends of clavicle Cardiovascular Coarctation of aorta or ventricular septal defect (10%–16%) Renal (38%) Horseshoe kidneys Unilateral renal aplasia Duplication ureters Gastrointestinal Telangiectasias Skin and Lymphatics Pigmented nevi (63%) Lymphadema (38%), generalized, due to hypoplasia superficial vessels; puffy hands and feet Nails Hypoplasia and malformation (66%) Skeletal Cubitus valgus (54%) Radial tilt of articular surface of trochlear Clinodactyly V Short metacarpals, usually IV (48%) Decreased carpal arch (mean angle 117) Deformities of medical tibial condyle Dermatoglyphics Increased total digital ridge count

X Short-Arm Deletions and Ovarian Failure Several different terminal deletions of the short arm of the X chromosome exist, showing varying amounts of

Increased distance between palmar triradii a and b Distal axial triradius in position t0 Percentage affected reflect tabulation of Simpson JL. Gonadal dysgenesis and abnormalities of the human sex chromosomes: current status of phenotypickaryotypic correlations. Birth Defects Orig Artic Ser 1975;11(4):23–59.

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OVARIAN FAILURE AS A RESULT OF HAPLOINSUFFICIENCY FOR THE X CHROMOSOME (X-MONOSOMY AND X-DELETIONS)

persisting Xp. Determining precise breakpoints and disrupted genes is necessary, but still not often pursued. Few studies have reexamined cases ascertained before the modern molecular era. Pooling morphologically similar terminal deletions has long shown that the most common breakpoint for terminal deletions is Xp11.2 ! 11.4. Again, the actual molecular breakpoint could and probably is different in literally every case. Overall, however, in 46,X,del(X) (p11) only proximal Xp remains; thus del(Xp) chromosome appears acrocentric or telocentric. Distal (telomeric) breakpoints are also reported including Xp21, 22.1, 22.2, and 22.3. Sequencing and analysis with polymorphic DNA markers are beginning to identify breakpoints. Approximately half of 46,X,del(Xp)(p11) cases show primary amenorrhea and gonadal dysgenesis. In one tabulation by the author 15 years ago, 12 of 27 reported del(X)(p11.2 ! 11.4) individuals menstruated spontaneously; however, menstruation was rarely normal. More recent calculations have not materially altered this conclusion. Fig. 1 shows our compilation of cases through 1999. To date, molecular analyses have yielded conclusions similar to those derived on the basis of phenotypickaryotypic correlations. Women with more distal deletions [del(X)(p.21.1 to p22.122)] menstruate more often, but many are still infertile or show secondary amenorrhea. Ovarian failure (menopause) is more likely to occur prematurely (younger than 35 or 40 years of age, depending on

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definition). Thus loci in regions Xp21, 22.1 or 22.2 are probably less pivotal for ovarian development than those nearer the centromere (Xp11). Deletion of only the most telomeric portion of Xp (Xp22.3 ! Xpter) does not result in amenorrhea. Zinn et al. [6] and Zinn and Ross [7] concluded only that the relatively large region Xp11.3 ! 22.1 was pivotal. Cases with an interstitial deletion will ideally help narrow the region of interest. [6,7] Familial cases of X short-arm deletions have been observed, namely mother and daughter having either the same terminal Xp-deletion or the same X/autosome translocations. Among 10 del(Xp) cases studied by James et al. [8] were two mother-daughter pairs. Thomas and Huson [9] reported 25 females with de novo deletions of Xp and four familial Xp deletions. Familial cases have involved deletions at Xp11 as well as Xp22 ! 12. Almost all reported 46,i(Xq) patients show streak gonads and short stature, and Turner stigmata have long been accepted as almost universal. Only rarely do 46,X,i (Xq) patients menstruate. The frequency of gonadal failure in 46,X,i(Xq) individuals is greater than in 46,X,del (Xp11) individuals, about half of whom menstruate or develop breasts. In turn, isochromosome for the X long arm [i(Xq)] differs from terminal deletion of Xp in that not just the terminal portion but all of the Xp is deleted. This finding further suggests the existence of gonadal determinants at several different locations on Xp. If so, a locus near the centromere could be deleted in i(Xq) yet retained in del(X)(p11).

FIG. 1 Schematic diagram of the X chromosome showing ovarian function as a function of nonmosaic terminal deletions.

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Candidate Genes for Xp Gonadal Determinants Several candidate genes for Xp gonadal determinants have been proposed. Zinn et al. [6] proposed ZFX, a DNA binding protein; a homologous gene exists on the Y (ZFY). Jones et al. [10] proposed Drosophila fat facets related X (DFRX), located on Xp11.4 and homologous to a locus on Yq11.2. DFRX (or USP9X) targets proteins for degradation by the ubiquitin pathway. USP9X is homologous to the Drosophila gene fat facets (faf). Both DFRX (USP9X) and ZFX escaped inactivation in two human de novo (X)(p11.2) deletions; however, after observing ovarian function in two cases despite haploinsufficiency, James et al. [11] concluded that DFRX was an unlikely candidate. On the other hand, neither of these cases was categorically normal clinically; therefore, a role for DFRX in gonadal development is not completely excluded. Studying an interstitial deletion of Xp11.2 ! 11.4, Zinn et al. [6] hypothesized three Xp candidate genes: DFRX (USP9X), ubiquitin activating enzyme E1 (UBE1, another ubiquitin pathway enzyme), and bone morphogenetic protein [BMP15, a member of the transforming growth factor-β (TGF-β) family of signaling compounds]. Mutations in BMP15 cause primary ovarian failure in X-linked recessive fashion in Inverdale sheep. BMP15 is structurally similar to GDF9 and probably interacts with GDF9 in vivo. BMP15 takes part in the regulation of follicular growth, maturation, and postovulatory process. Mutations of BMP15 have been found responsible for POF in Caucasian, Indian and Chinese patients; however, its mutation frequencies varied from 1.5% to 15%. [12] Most of these variants are heterozygous, such as p.R329C in the coding region and p. R68W, p. R138H in the pro-region, indicating that sufficient dose of BMP15 is necessary for normal ovarian reserve. [13] Other candidate X-ovarian genes are reviewed by Bione and Toniolo [14], Zinn [15], and Zinn and Ross [7]. In addition, many other candidate genes can be deduced from homologues in the mouse [see Simpson and Rajkovic [1] and Matzuk and Lamb [16]]. The latter provided an overview of murine genes causing germ-cell abnormalities in males and females; however, in the mouse most of these genes are autosomal. Thus perhaps better candidate genes could be proposed for the disorders of human XX gonadal dysgenesis to POF. Irrespective, determining which human homologues of these mouse genes are pertinent will be the future direction.

X Long-Arm Deletions Deletions of the X long arm (Xq) vary in composition, as do those involving the X short arm. The most extensive deletions originate at Xq13 and are associated invariably with primary amenorrhea, lack of breast development, and complete ovarian failure. [1,11,17] Xq13 must be a

pivotal region for ovarian maintenance. That this region contains the human X-inactivation center (XIC) may or may not be the explanation. The abnormal phenotype associated with del(Xq13) could reflect perturbation of XIC and loss of the gene product Xist rather than loss of an ovarian gene per se. Irrespective, key loci must lie no more distal than proximal Xq21, given that menstruation occurs in deletions of breakpoints Xq21 or beyond (see Fig. 1). Menstruating del(X)(q21) women might have retained a region containing an ovarian maintenance gene, whereas del(X)(q13 or 21) cases with primary amenorrhea could have lost such a locus. In more distal Xq deletions, the phenotype is usually not primary amenorrhea but rather POF [1]. Distal Xq is thus less pivotal for ovarian maintenance than proximal Xq although the former still contains regions important for ovarian maintenance. Although there is no stepwise demarcation into discrete breakpoint regions, it is heuristically useful to stratify terminal deletions in this fashion: Xq13 ! 21, Xq22 ! 25, and Xq26 ! 28. Ogata and Matsuo [17] correlated ovarian function using such stratification and arrived at conclusions similar to our own. Molecular mapping of the regions of Xq integral for ovarian development has begun, but to date views on locations and the nature of Xq ovarian determinants have not changed substantively from those derived on the basis of phenotypic-karyotopic correlations. Sala et al. [18] studied seven X/autosome translocations involving Xq21 ! 22, five of which had primary amenorrhea. A region of Xq spanning 15 mb encompassed breakpoints in all seven cases. Breakpoints in four other X-autosome translocations studied by Philippe et al. [19] were localized to the same region, extending from DXS233 to DXS1171 (most of Xq21). [20] If the breakpoints associated with ovarian failure truly spanned the entire Xq21 region, it would be unlikely that only a single gene in this region is responsible for ovarian failure. Alternatively, ovarian failure associated with these translocations could be the result not of perturbation of any specific gene but rather of generalized cytologic (meiotic) instability. Incidentally, the official nomenclature designates genes causing POF as POF1 and POF2, based on presumptive order of discovery. Thus the Xq23 ! Xq27 region is POFl, whereas Xq13 ! Xq21 region is officially POF2. Distal Xq deletions may be familial, such as del(Xp). Some of these familial aggregates are derivative of Xq/ autosome translocations, but aggregating with terminal deletions exist as well [19]. Ovarian function may vary among different family members having the same deletion.

Candidate Genes for Xq Gonadal Determinants A popular candidate for genes causing ovarian failure on the X long arm is the human homolog of the Drosophila melanogaster gene diaphanous (dia). Drosophila dia is a

V. HUMAN OVARIAN PATHOPHYSIOLOGY: SELECT ASPECTS

OVARIAN FAILURE AS A RESULT OF MENDELIAN CAUSES

member of a family of proteins that establish cell polarity, govern cytokinesis, and organize the cytoskeleton mutation of the dia gene, which causes sterility in male and female flies. Sequence comparisons between dia and the human EST (expressed sequence tag) DRE25 show significant homology. In turn, DRE25 maps to human Xq21, a region we have already noted to be important for ovarian maintenance. Studying an Xq21/autosome translocation originally reported by Philippe et al. [19], Sala et al. [18] found DRE25 to be disrupted in women with ovarian failure. Another candidate gene is XPNPEP2, which encodes the Xaa-Pro aminopeptidase (metalloprotease) that hydrolyzes proline bonds. XPNPEP2 is ubiquitously expressed in many tissues and can influence a host of biologically active peptides that could include regulations of ovarian function. XPNPEP2 was disrupted in an Xq translocation that involved Xq25 [14–16], and several other Xq candidate genes were considered. Several other genes on the X long arm have also been associated with ovarian failure. • Progesterone receptor membrane component 1 (PGRMC1) mediates progesterone’s anti-apoptotic effects on granulosa cells. A missense mutation (p. H165R) in the PGRMC1 gene was suggested to result in POF through impaired activation of the microsomal cytochrome P450, and increased apoptosis of ovarian cells [21]. • The androgen receptor (AR) gene encodes androgen receptor and is involved in sex differentiation and reproduction. Female Ar/ mice showed subfertility and decreased follicle count with aberrant ovarian gene expression. In addition, the CAG repeat length in exon 1 of the AR gene is presumed to be associated with POF, but still remains controversial. • FOXO4, belonging to O class of winged helix/forkhead transcription factor family (FOXO), localizes to granulosa cells in mice and human. This gene involves in PI3K/Akt/Cdkn1b molecular pathway, which suggests a functional role during ovarian physiology. • POF, 1B (POF1B), which is considered to escape X-chromosome inactivation and critical for ovarian function is located within POF1 region (Xq23 ! Xq27). In addition, POF1B binds to nonmuscle actin filament and regulates meiotic cell division, as well as epithelial monolayers organization. • Other candidate genes on X long arm include diaphanous-related formin2 (DIAPH2, Xq22), X-prolyl aminopeptidase (aminopeptidase P) 2, membranebound (XPNPEP2, Xq25), dachshund family transcription factor 2 (DACH2, Xq21.3), choroideremia (CHM, Xq21.1), collagen, type IV, alpha 6 (COL4A6, Xq22.3), and nuclear RNA export factor 5 (NXF5, Xq22.1) et al. [12].

465

OVARIAN FAILURE AS A RESULT OF MENDELIAN CAUSES Phenotypic females with normal chromosomal complements (46,XX) may have gonadal failure and display ovarian failure identical to that of 45,X gonadal dysgenesis (Turner syndrome). Applying the general term XX gonadal dysgenesis implies that a coexisting 45,X line does not exist. A wide spectrum of disorders falls under this rubric (Table 2). In this section we discuss those disorders of ovarian failure whose etiology is either known to be caused by autosomal genes or which seems likely to be caused by such.

XX Gonadal Dysgenesis Without Somatic Anomalies This category implies a diagnosis of exclusion for those 46,XX gonadal dysgenesis cases not associated with somatic anomalies. Affected individuals are normal in stature (mean height, 165 cm), and somatic features of TABLE 2

The Clinical Spectrum of XX Gonadal Dysgenesis

XX gonadal dysgenesis without somatic anomalies XX gonadal dysgenesis with neurosensory deafness (Perrault syndrome) XX gonadal dysgenesis with cerebellar ataxia (hetereogeneous) XX gonadal dysgenesis in malformation syndrome (Table 3) XX gonadal dysgenesis as one component of pleiotropic Mendelian disorders (Table 4) LH receptor mutations (LHR) FSH receptor mutations (FSHR) Blepharophimosis-ptosis-epicanthus (FOXL2) Germ cell absence in both sexes (46,XX) Adrenal and ovarian biosynthetic defects Aromatase 17a-hydroxylase (CYP17) Agonadia (46,XX cases) Inborn errors of metabolism Galactosemia Carbohydrate-deficient glycoprotein (phosphomannomutase deficiency, PMM2) Dynamic mutations (triplet repeat) Fragile X (FRAXA) Myotonic dystrophy (uncommon cause) Ovarian-specific autoimmunity Polyglandular autoimmune syndrome (premature ovarian failure) Autosomal trisomies Trisomy 13 Trisomy 18

V. HUMAN OVARIAN PATHOPHYSIOLOGY: SELECT ASPECTS

466

28. GERM CELL FAILURE AND OVARIAN RESISTANCE: HUMAN GENES AND DISORDERS

Turner stigmata are absent. The known candidate genes on autosomal chromosomes have been categorized according to their biological functions. Genes regulating oogenesis and follicular development include BMP15, forehead box L2 (FOXL2), spermatogenesis and oogenesis specific basic helix–loop–helix 1 (SOHLH1), newborn ovary homeobox gene (NOBOX), folliculogenesis specific bHLH transcription factor (FIGLA), growth differentiation factor (GDF9); genes affecting hormone synthesis, such as follicle-stimulating hormone receptor (FSHR), luteinizing hormone receptor (LHR), cytochrome P450 Family 17 subfamily A member 1 (CYP17) and cytochrome P450 family 17 subfamily A member 1 (CYP19); genes involved in meiosis and DNA damage repair, such as stromal antigen 3 (STAG3), ATP-dependent DNA helicase homolog (HFM1), minichromosome maintenance complex component 8 and 9 (MCM8, MCM9), Cockayne syndrome group B (CSB)—PiggyBac transposable element derived 3 (PGBD3) (Table 3). Online Mendelian inheritance in man (OMIM) has nominated 14 genes as causative genes as POF1 to POF13, 5 of which are located on the X chromosome, FMR1 (POF1), DIAPH2 (POF2A), POF1B (POF2B), FOXL2 (POF3), and BMP15 (POF4), the remaining 9 genes on autosomal. These 9 genes, NOBOX, FIGLA, nuclear receptor subfamily 5, group A, member 1 TABLE 3

Location

Case

Control

Ethnicity

HFM1

1p22.2

2(family)

316

Chinese

69

316

Chinese

1p31

Amino acid change

c.T2651G

p.I884S

c.G2206A

p.G736S

c.3929_3930 del insG

p.P1310Rfs*41

None

2(family)

135

Colombian

c.2355 + 1G>A

None

94

Caucasian

TGFBR3

1p33–p32

112

110

Chinese

2 (1.8%)

p.I743_K785del

c.1370A>G

p.E458G

c.2467C>T

p.P824L

54

41

New Zealand

133

200

Indian

100

100

Chinese

None

Caucasian

None

82 1p36.23–p35.1

2(2.9%)

Sequence variation

Caucasian Senegalese

95

1p36.1–p35

Mutation ratea

36

1p31.1

WNT4

Homeobox transcription factor NOBOX is highly expressed in oocytes and known to play a critical role in early folliculogenesis. In female mice, disruption of the Nobox gene causes acceleration of postnatal oocyte loss, transition abolishment from primordial to growing follicles, and replacement of follicles by fibrous tissue, which is consistent with a human POF phenotype. In 2007, Qin et al. [22] demonstrated, for the first time, that a missense mutation (p.R355H) in NOBOX was responsible for human POF. The mutation disrupted NOBOX homeodomain binding to NOBOX DNA-binding element (NBE) and had a dominant negative effect. Subsequently, Bouilly et al. [23,24] sequenced NOBOX gene in POF cohorts in 2011 and 2015. They identified the frequencies of the novel loss-of-function mutations in Caucasian and African ancestry to be 6.2% and 5.6%, respectively. Therefore, NOBOX may be the most common autosomal gene most frequently implicated in POF.

41

LHX8

GPR3

NOBOX (POF5)

Variants Identified in Candidate Genes on Autosomes for Sporadic and Familial Patients With POF

Gene

MSH4

(NR5A1), STAG3, HFM1, MCM8, PGBD3, synaptonemal complex central element 1 (SYCE1), MutS homolog 5 (MSH5) are further discussed in detail.

None 1 (0.8%)

c.2323C>T

55

100

Tunisian

None

145

200

Chinese

None

p.P775S

V. HUMAN OVARIAN PATHOPHYSIOLOGY: SELECT ASPECTS

FC

Mechanism

Yes

Led to the ablation of the Walker B motif and inactivated MSH4.

467

OVARIAN FAILURE AS A RESULT OF MENDELIAN CAUSES

TABLE 3

Variants Identified in Candidate Genes on Autosomes for Sporadic and Familial Patients With POF—cont’d

Gene

Location

Case

EXO1

1q43

186

FIGLA

2p13.3

100

219 FSHR

2p21–p16

Control

Ethnicity Chinese

304

230

36

Chinese

Indian

2 (0.9%)

Amino acid change

None

None

c.15-36del

p.G6fsX66

c.419-421del

p.140delN

20

93

Germany

None

20

44

Argentine

None

16

236

Singapore Chinese

None

49

51

UK

None

50

50

New Zealand

None

50

50

Indian

73

35

Chinese

15

3

Japanese

15

42

Brazilian

1094

Denmark

1162

Switzerland

540

Singapore Chinese

1976

Finnish

19 (0.96%)

c.566C> T

p.A189V

52

Finnish

22(42.3%)

c.566C> T

p.A189V

New Zealand Slovenian

None None

43

INHA

2q33–q36

50

50

Indian

133

200

Indian

1(0.09%)

c.566C> T

2 (1.5%)

c.525C> G

p.H175O

c.769G>A

p.A257T

150

New Zealand Slovenian

None

Chinese

None

Indian

None

Indian

None

70

30

100

New Zealand Slovenian Unknown

9 (11.2%)

p.S92 N

43

120

Yes

Disrupted binding to the TCF3 HLH domain

Yes

Dramatic reduction of binding capacity and signal transduction

p.A189V

c.275G>A

Indian

50

Truncated protein

None

100

80

Mechanism

None

80

118

FC

c.649 + 8A> G None

2cen–q13

3q23

2 (2.0%)

Sequence variation

Southern Brazilian

INHBB

FOXL2

Mutation rate

2 (2.9%)

c.898-927del

p.A221_A230del

c.1009T>A

p.Y258N

None Continued

V. HUMAN OVARIAN PATHOPHYSIOLOGY: SELECT ASPECTS

468 TABLE 3

28. GERM CELL FAILURE AND OVARIAN RESISTANCE: HUMAN GENES AND DISORDERS

Variants Identified in Candidate Genes on Autosomes for Sporadic and Familial Patients With POF—cont’d Sequence variation

Amino acid change

Caucasian African American Hispanic

c.798C>T

Sense

Chinese

None

200

Chinese

None

101

106

French

None

5q11.2

80

80

Chinese

c.270C>G

5q31.1

100

100

Chinese

None

20

93

Germany

None

100

96

Chinese

1 (1.0%)

c.712A>G

p.T238A

61

60

American

1 (1.6%)

c.307C>T

p.P103S

203

54

Caucasian African Asian

1 (0.5%)

c.557C>A

p.S186Y

38

51

New Zealand

127

220

Indian

Gene

Location

Case

Control

Ethnicity

KIT

4q11–q12

40

10

HELQ

4q21.23

192

SKP2

5p13

200

PRLR

5p13.2

FST GDF9

15

3

Japanese

Mutation rate

Yes

Decreased mature protein production, weaker Smad1/5/8 phosphorylation in COV434 cells and decreased granulosa cell proliferation

Yes

Decreased mature protein production, weaker Smad1/5/8 phosphorylation in COV434 cells and decreased granulosa cell proliferation

Yes

Impaired DNA homologous recombination repair

None 6 (4.7%)

c.199A>C

p.K67E

c.646G>A

p.V216 M

None

6p21.31

115

149

Chinese

1 (0.9%)

c.37C>A

p.P13T

MSH5

6p21.3

41

36

Caucasian Senegalese

2 (4.9%)

g.2547C>T

p.P29S

2(family)

400

Chinese

c.1459G>T

p.D487Y

200

400

Chinese

114

100

Chinese

c.71C>A

p.P24H

c.140C>T

p.P47L

c.184G>A

p.D62N

c.1652C>T

p.S551F

6q21

Mechanism

Sense

POU5F1

FOXO3

FC

15 (13.2%)

V. HUMAN OVARIAN PATHOPHYSIOLOGY: SELECT ASPECTS

469

OVARIAN FAILURE AS A RESULT OF MENDELIAN CAUSES

TABLE 3 Gene

Variants Identified in Candidate Genes on Autosomes for Sporadic and Familial Patients With POF—cont’d Location

Case

Control

302

22

50 90

MCM9

INHBA

6q22.31

7p15–p13

60

Ethnicity

Mutation rate

Sequence variation

Amino acid change

c.1697C>G

p.G566A

c.280C> T

p.L94F

c.1021G>A

p.A341T

c.1156C>T

p.L386F

Italy North America France Germany South Korea

4 (1.3%)

French

3 (6.0%)

c.1778A>C

p.Y593S

New Zealand, Slovenia

2 (2.2%)

c.1262C>T

p.S421L

c.1517G>A

p.R506H

2(family)

Kurdish

151

White, Hispanic, Indian, (mixed) African American

5(3.3%)

c.1651C>T

p.Q551*

c.1784C>G

p.T595R

c.2011G>T

p.E670*

c.2422G>A

p.V808I

None

43

New Zealand Slovenian

None

Palestinian

c.968delC

p.F187 fs*7

c.271G>T

p.G91 T

c.331G>A

p.G111R

c.349C> T

p.R117W

c.1112A>C

p.K371 T

c.1856C>T

p.P619L

c.271G>T

p.G91 W

c.349C> T

p.R117W

c.1025G>C

p.S342 T

c.1048G>T

p.V350 L

c.907C> T

p.R303X

5(family)

NOBOX

7q35

213

178

362

362

Caucasian or African

Caucasian Senegalese Bantu

200

200

Chinese

96

278

Caucasian

Yes

No effect on transactivation activity of FHRE-luc

Anormal alternative splicing and truncated forms of MCM9

Indian

7q22.1

Mechanism

c.1732 + 2T>C

80

STAG3

FC

12 (5.6%)

12 (6.2%)

Yes

Loss of function of the pathogenic mutants due to their inability to binding with DNA.

Yes

Compromised the ability to bind to and transactivate GDF9 promoter

Yes

Disrupted binding to NOBOX DNAbinding element NBE with a dominant negative effect

None 1(1.0%)

c.1064G>A

p.R355H

V. HUMAN OVARIAN PATHOPHYSIOLOGY: SELECT ASPECTS

Continued

470 TABLE 3 Gene

28. GERM CELL FAILURE AND OVARIAN RESISTANCE: HUMAN GENES AND DISORDERS

Variants Identified in Candidate Genes on Autosomes for Sporadic and Familial Patients With POF—cont’d Location

Case

Control

Ethnicity

30

20

Japanese

96

211

Chinese

252

Israeli Muslim Arab

SPIDR

8q11.21

2(family)

NR5A1

9q33

180

French

Amino acid change

FC

Mechanism

c.567delG

p.T190Hfs*13

Yes

Defects in NOBOX transcriptional activation

c.839G>A

p.W280*

Yes

Impaired repair ability for DSBs

c.162C>A

p.S54R

Yes

c.593C>T

p.P198L

No effect on DNA binding capacity and transcriptional activity

None 1(1%)

3 (1.7%)

50

Tunisian

1 (1.8%)

c.763C>T

p.R255C

Yes

Marked decrease in transactivation on the Cyp11a1 and Amh promoters

384

400

Chinese

1(0.26%)

c.13T>G

p.Y5D

Yes

Impaired transcriptional activation on Amh, Inhibin-a, Cyp11a1 and Cyp19a1 gene

55

100

Tunisian

Yes

Severe reduction in transactivation of CYP11A1 and CYP19A1

Yes

Impaired GCs differentiation, and oocyte-GCs interaction

Asian Caucasian Mediterranean

28

9q34.3

Sequence variation

26

356

SOHLH1

Mutation rate

400/479

561

600

None 5 (1.4%)

Roman Indian Senegalese African

2 (2.3%)

Chinese Serbian

2 (0.36%)

c.407C>T

p.P136L

c.574G>T + c.575C>T

p.A192F/S + A192V

c.593C>T

p.P198L

c.938G>A

p.R313H

c.691-699del

p.L231_L233del

c.368G>C/ c.386C>T

p.G123A/P129L

c.950C>T

p.S317F

c.1126G>A

p.E376K

CSBPGBD3

10q11.23

family

PTEN

10q23.3

148

378

Chinese

None

20

20

Japanese

None

Chinese

NANOS1

10q26.11

100

200

Chinese

None

SYCE1

10q26.3

2(family)

90

Arabian

c.613C>T

p.Q205*

WT1

11p13

384

384

Chinese

c.376C>T

p.P126S

c.1109G>A

p.R370H

CDKN1B

12p13.1–p12

200

200

Chinese

87

263

Tunisian Colombian

2(0.5%)

None 1 (1.5%)

c.356T>C

p.I119T

V. HUMAN OVARIAN PATHOPHYSIOLOGY: SELECT ASPECTS

TABLE 3 Gene

AMHR2

Variants Identified in Candidate Genes on Autosomes for Sporadic and Familial Patients With POF—cont’d Location

12q13

Mutation rate

Sequence variation

Case

Control

Ethnicity

124

100

Chinese

None

16

12

Japanese

None

96

Chinese

Chinese

None None

PRIM1

12q13.3

192

KITLG

12q22

40

70

Caucasian

SOHLH2

13q13.3

561

600

Chinese Serbian

1

8 (1.4%)

c.50C> A

Amino acid change

FC

Mechanism

p.A17E

Yes

No severe defects in transducting the AMH signal

p.R153W

Yes

Decreases the stability of NANOS3 and result in hypomorph

c.235G>A

p.E79K

c.314A>G

p.E105G

c.360A>T

p.L120F

c.610C> T

p.L204F

c.961A>C

p.T321P

c.251C> T

p.P84L

FOXO1

13q14.1

90

POLG

15q25

57

British

None

38

Italian

None

201

Caucasian black Hispanic Asian

1 (0.5%)

c.2857C>T

p.R953C

1 (1.0%)

c.275G>A

p.G92E

60

New Zealand, Slovenia

1 (1.1%)

NOG

17q22

100

43

French

AMH

19p13.3

16

12

Japanese

None

NANOS3

19p13.13

168

63

Chinese Caucasian

None

100

200

Chinese

1(1%)

c.457C> T

NANOS2

19q13.32

100

200

Chinese

None

MCM8

20p12.3

3(family)

200

Saudi Arabian

c.446C> G

p.P149R

Yes

Inhibited recruitment of MCM8 to sites of DNA damage

192

312

Chinese

c.A950T

p.H317L

Yes

c.A1802G

p.H601R

Impaired repair ability for DSBs

155

SALL4

20q13.2

100

300

2(1%)

White, Hispanic, Indian, (mixed) African American

1

c.G1443A

p.R445Q

Chinese

2 (2%)

c.541G>A

p.V181 M

c.2449A>G

p.T817A

SPO11

20q13.31

41

36

Caucasian Senegalese

None

DMC1

22q13.1

192

400

Chinese

None

41

36

Caucasian Senegalese

None

a

Only refer to novel missense, frameshift and nonsense mutations identified in patients with sporadic POF.

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28. GERM CELL FAILURE AND OVARIAN RESISTANCE: HUMAN GENES AND DISORDERS

FIGLA (POF6) FIGLA is a germ-cell specific, basic helix-loop-helix (bHLH) transcription factor, regulating the formation of primordial follicle, and coordinating expression of zona pellucida genes. FIGLA knockout mice cannot form primordial follicles and lose oocytes rapidly after birth, whereas male gonads are unaffected. In 2008, Zhao et al. [25] reported two plausible mutations in FIGLA among 100 Chinese women with POF. One is deletion mutation p.G6fsX66, which leads to haploinsufficiency; the other one is deletion mutation p.140 delN, which results in disruption of FIGLA blinding to the TCF3 helix-loop-helix (HLH) domain and having dominant negative effect. NR5A1 (POF7) NR5A1 encodes steroidogenic factor 1 (SF1). As a nuclear receptor, NR5A1 is involved in adrenal and gonadal development, steroidogenesis, and reproduction. Knockout of Nr5a1 in granulosa cells causes infertility and hypoplastic ovaries with reduction of follicles and absence of corpora lutea. In human, NR5A1 is associated with 46,XY disorder of sex development (DSD) and POF. In 2012, Janse et al. [26] sequenced NR5A1 in 386 wellphenotyped women with POF. The mutation frequency was 1.4%; however, bioinformatic prediction showed that the pathogenesis of NRA51 mutation was lower than intermediate level. In 2013, Voican et al. [27] screened NR5A1 in 180 patients with idiopathic POF, and identified four missense variants in three patients. Further functional analysis found no deleterious effects of mutant protein on DNA binding capacity and transcriptional activity compared with wild-type NR5A1. Another novel mutation (p.Y5D) in Chinese was presumed to result in haploinsufficiency and responsible for POF [28]. STAG3 (POF8) The STAG3 gene encodes a subunit of cohesion ring, which plays a vital role in the formation of the synaptonemal complex, participating in the proper pairing and segregation of chromosomes during meiosis, and being indispensable for fertility. Inactivation of Stag3 in mice causes severe ovarian dysgenesis, lacking of oocytes and follicles. In 2014, the homozygous deletion c.968delC was found in a large family with inherited POF and proved causative [29]. Recently, He et al. [30] identified a homozygous donor splice site mutation in the STAG3 gene in a consanguineous Han Chinese family presenting with primary amenorrhea. HFM1 (POF9) HFM1 is a meiotic gene encoding DNA helicase, which is necessary for normal progression of homologous recombination and proper synapsis between homologous

chromosomes. Hfm1/ female mice appeared to have eliminated most of crossovers during meiosis and showed significant reduction in ovary size, follicles, and corpora lutea numbers. In 2014, Wang et al. [31] identified compound heterozygous mutations (c.1686-1G.C and p. I884S) of HFM1 gene in two affected Chinese sisters, which led to autosomal recessive POI. Another compound heterozygous mutation (p.G736S and p.P1310R fs*41) was found through screening of HFM1 in 69 Chinese women with sporadic POF; none of these variants were present in 316 matched controls. MCM8 (POF10) MCM8 belongs to the evolutionarily conserved MCMs protein family and is responsible for homologous recombination and double-stranded DNA break (DSB) repairs. MCM8 is not only involved in the assembly of the prereplication complex during G1 phase, but also participates in homologous recombination during meiosis and DSB repair by dimerizing with MCM9. Mcm8 knockout mice showed small gonads and early block of follicle development. Rajkovic and colleagues [32] identified a homozygous mutation p.P149R in a consanguineous family with three affected daughters with primary amenorrhea via single nucleotide polymorphism analysis and whole exome sequencing. The variant inhibits recruitment of mutant MCM8 to sites of DNA damage, and finally endocrine dysfunction and genomic instability. In 2016, Dou et al. [33] screened the coding regions of MCM8 in 192 Chinese patients with sporadic POF and identified two novel heterozygous mutation p. H317L and p. H601R. Further functional study proved that mutant p. H317L impaired its repair ability for DSBs. CSB-PGBD3 (POF11) CSB-PGBD3 is a fusion gene that encodes a protein participating in transcription-coupled DNA repair (TCR) of DNA damage. Qin et al. [34] identified a heterozygous p.G746D mutation in CSB-PGBD3 through whole exome sequencing in a nonconsanguineous Chinese family with POF. Another missense mutation p.V1056I and one nonsense mutation (p.E215X) in CSB-PGBD3 were found in 432 sporadic cases. Furthermore, the truncated protein E215X manifested loss of function; mutant p. G746D, p.V1056I, and p.E215X appeared delayed or nonresponsive to DNA damage caused by laser/UV/H2O2. SYCE1 (POF12) SYCE1 encodes synaptonemal complex central element protein 1, a component of the synaptonemal complex that links paired homologous chromosomes. Disruption of Syce1 gene in female mouse appeared infertility and completely lacking of follicles, suggesting defects in meiosis and oogenesis. Different hemizygous deletions were found in Caucasian and Chinese patients

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with POF through CNV analysis. Vries et al. [35] identified a protein-truncating homozygous mutation (p.Q205*) in the SYCE1 gene in sisters with POF from consanguineous family. This study further elucidated the importance of SYCE1 involved in ovarian function. MSH5 (POF13) MSH5 belongs to the DNA mismatch repair gene family, exerting essential functions for normal chromosome synapsis during zygotene, and meiotic recombination. Disruption of Msh5 in female mice resulted in sterility, ovarian degeneration, and progressive loss of oocytes due to meiotic failure. A heterozygous mutation p.P29S in MSH5 was found in 2 of 41 Caucasian patients with POF, whereas lacking functional confirmation [36]. Recently, Guo et al. [37] found that in a Chinese pedigree two sisters with POF carried homozygous missense mutation p.D487Y in the MSH5. The homologous mutation in mice resulted in atrophic ovaries without oocytes. in vitro functional studies revealed that mutant MSH5 impaired DNA homologous recombination repair. This study confirmed that when DNA damage repair gene was perturbed, it might lead to non-syndromic POF.

Perrault Syndrome (XX Gonadal Dysgenesis With Neurosensory Deafness) XX gonadal dysgenesis associated with neurosensory deafness has long been recognized and called Perrault syndrome. Perrault syndrome is genetically heterogeneous and inherited in autosomal recessive fashion. Multiple candidate genes have been identified by whole exome sequencing, including hydroxysteroid (17-beta) dehydrogenase 4 (HSD17B4), histidyl-tRNA synthetase 2, mitochondrial (HARS2), leucyl-tRNA synthetase 2, mitochondrial (LARS2), caseinolytic mitochondrial matrix peptidase proteolytic subunit (CLPP), and Chromosome 10 open reading frame 2 (C10orf2). HSD17B4 was the first causative gene identified in Perrault syndrome. It is a multifunctional peroxisomal enzyme, also known as D-bifunctional protein (DBP), and participates in fatty acid β-oxidation and steroid metabolism. Unlike other causative genes for Perrault syndrome, DBP deficiency caused by mutations in HSD17B4 usually results in a severe phenotype within the first 2 years of life. Compound heterozygous mutations p.Y217C and p.Y568X were found in two sisters from a mixed European ancestry family. However, DBP deficiency resulting from these variants is relatively mild, which may allow survival to the age of puberty and be more likely to induce ovarian dysgenesis [38]. Nuclear genes encoding mitochondrial proteins are essential for the normal functions of mitochondria.

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Several are causative for Perrault syndrome. Mitochondrial tRNA synthetase, encoded by HARS2 and LARS2, is responsible for supplementing mitochondrial tRNAs with their cognate amino acids. Pierce et al. [39] identified compound heterozygous mutations p.L200V and p.V368L in HARS2 in a nonconsanguineous family from a mixed European ancestry. Later, they detected in LARS2 [40] homozygous mutation p.T522N in a consanguineous Palestinian family and compound heterozygous mutation c.1077delT and p.T629M in a nonconsanguineous Slovenian family. Perturbation in mitochondrial protein translation caused by genetic factors may result in tissue-specific apoptosis, further implicating in hearing loss and ovarian dysfunction. CLPP is a mitochondrial ATP-dependent chambered protease, accounting for protein degradation and mitochondrial homeostasis. Homozygous mutations p.T145P, p.C147S, and c.270 + 4A > G in CLPP were observed in affected individuals from three consanguineous Pakistani families with Perrault syndrome (including ovarian failure). [41] C10orf2, which encodes a primase-helicase, is required for mitochondrial DNA (mtDNA) replication and is also responsible for Perrault syndrome. Compound heterozygous mutations p.R391H plus p.N585S and p.W441G plus p.V507I in C10orf2 gene were found in a Japanese family and an English ancestry family, respectively.[42]

Cerebellar Ataxia and XX Gonadal Dysgenesis XX gonadal dysgenesis can be found in association with a heterogeneous group of cerebellar ataxias. The hereditary ataxias are confusing nosologically, principally because of ill-defined diagnostic criteria and lack of direct access to the cerebellum. Forms of ataxia characterized by hypogonadotropic hypogonadism also exist but are not relevant here. The association between hypergonadotropic hypogonadism and ataxia was first reported by Skre et al. [43]. In one family, a 16-year-old girl was affected. In the other family, three sisters were affected. In the one sporadic case and in one of the three affected sibs, ataxia was manifested shortly after birth; in the two other sibs, age of onset occurred later in childhood. Cataracts were present in all individuals described by Skre et al. [43]. Hypergonadotropic hypogonadism and ataxia were subsequently reported by De Michele et al. [44], Linssen et al. [45], Gottschalk et al. [46], Fryns et al. [47], Nishi et al. [48], and Amor et al. [49]. In these various reports, the clinical features of ataxia and other neurologic abnormalities varied (e.g., being either progressive versus nonprogressive) (see Simpson and Elias for further discussion).

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XX Gonadal Dysgenesis in Other Unique Malformation Syndromes Among other pleiotropic genes causing XX gonadal dysgenesis and distinctive somatic features are several that have been reported in only a single family. Table 4 lists these syndromes: XX gonadal dysgenesis, microcephaly, and arachnodactyly; XX gonadal dysgenesis, cardiomyopathy, blepharoptosis, and broad nasal bridge; XX gonadal dysgenesis and epibulbar dermoid; XX gonadal dysgenesis, short stature, and metabolic acidosis. If Mendelian, these disorders are presumably autosomal recessive; however, subtle chromosomal rearrangements cannot be excluded. In these syndromes and others that doubtless will be reported in the future, the key underlying biologic question is whether the ostensibly pleiotropic gene(s) causes both somatic anomalies and ovarian failure. If so, do the purported genes play roles in normal ovarian differentiation, or does their perturbation merely cause ovarian failure secondary to generalized somatic disturbance? Alternatively, could the somatic and gonadal phenotypes merely reflect closely linked genes (i.e., a contiguous gene syndrome)?

Pleiotropic Mendelian Disorders Showing Ovarian Failure Primary ovarian failure is observed in several well established and not necessarily uncommon Mendelian disorders. These are well known to geneticists and relevant to clinical specialists. All are characterized by distinct somatic features (Table 5). Of special note are Denys–Drash syndrome and Frasier syndrome, both of which are caused by mutations in the WT1 (Wilms’ tumor-1) gene. WT1 mutations can cause either 46,XY gender reversal or 46,XY genital ambiguity. One 46,XX individual with Frasier syndrome has been reported. This woman manifested not only the renal TABLE 4

Malformation Syndromes With 46,XX Gonadal Dysgenesis

Somatic features

Etiology (gene)

Cerebellar ataxia, sensorineural deafness, other somatic features

Autosomal recessive, heterogeneous

Microcephaly, arachnodactyly

Autosomal recessive

Epibulbar dermoids

Autosomal recessive

Short stature and metabolic acidosis

Autosomal recessive

Denys–Drash/Frasier syndrome

Autosomal dominant

Dilated cardiomyopathy, mental retardation, bleparoptosis (Malouf syndrome)

Autosomal recessive

parenchymal disease characteristic of Frasier syndrome, but also primary amenorrhea and ovarian failure. Gonadal failure in 46,XX Frasier syndrome could easily pass unappreciated if the primary amenorrhea were assumed secondary to azotemia.

Germ-Cell Failure in Male (46,XY) and Female (46,XX) Sibs In several sibships, male (46,XY) and female (46,XX) sibs have each shown germ-cell failure. Affected 46,XX females showed streak gonads, whereas 46,XY males showed germ-cell aplasia (Sertoli-cell-only syndrome). In two families, parents were consanguineous, and in each there were no somatic anomalies. In three other families, characteristic somatic anomalies suggested distinct entities. Hamet et al. [50] observed germ-cell failure, hypertension, and deafness; Al-Awadi et al. [51] found germ-cell failure and alopecia; Mikati et al. [52] reported germ-cell failure, microcephaly, and short stature. In each of these families, a single autosomal gene is presumed to deleteriously affect germ-cell development in both sexes. This gene(s) could act either at a specific site common to early germ-cell development or exert its effect through meiotic breakdown. Elucidating such genes could help explain normal germ cell development. Proliferation of germ cells (POG, now known as Fanconi anemia complementation group L, FANCL) was shown to be disrupted in germ cell deficient mice and presumably responsible for the observed phenotype both in males and females.[53] POG is one of the few genes known to specifically affect primordial germ-cell proliferation at the time of their entry into the developing genital ridge. Several other genes are proven to affect either PGC specification, namely beta-interferon gene positiveregulatory domain I binding factor (BLIMP1), homobox protein NANOG (NANOG), POU domain, class 5, transcription factor 1 (POU5F1) and nanos homolog 3 (NANOS3), others affect survival, proliferation, and migration of germ cells, namely tyrosine kinase receptor (KIT) and its ligand (KITL), deleted in azoospermia-like (DAZL), and DEAD-Box Helicase (DDX4).[54] NANOS3, a member of NANOS family, is required for PGC development and maintenance via suppression of apoptosis. NANOS3 is expressed in germ cells both in human ovary and testis, and localizes to the nucleus with co-expression with known germ cell proteins, BLIMP1, DDX4, and STELLA. NANOS3 plays an important role in human oogenesis and spermatogenesis. Nanos3/ mice show decreased gonad size and infertility in male and female. A potential heterozygous mutation (p.R153W) was identified in 100 Chinese patients with POF.[55] Another homozygous mutation (p.E120K) was also found in two Brazilian sisters with primary

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TABLE 5

Autosomal-Recessive Disorders in Which the Phenotype Is Predominately Somatic, But Includes Ovarian Failure [3]

Disorder

Causative gene

Ataxia–telangiectasia

Somatic features

Ovarian anomalies

Etiology

ATM

Cerebella ataxia, multiple telangiectasias (eyes, ears, flexa surface of extremities), immunodeficiency, chromosomal breakage, malignancy, X-ray hypersensitivity

“Complete absence of ovaries,” “absence of primary follicles”

Autosomal recessive

Carbohydrate-deficient glycoprotein syndrome, Type 1

PMM2

Neurologic abnormalities (e.g., unscheduled eye movements), ataxia, hypotonial/hyporeflexia strokes, joint cartractures

Ovarian failure (hypogonadism)

Autosomal recessive

Cockayne syndrome

ERCC6

Dwarfism, microcephaly, mental retardation, pigmentary retinopathy and photosensitivity, premature senility. Sensitivity to ultraviolet light

Ovarian atrophy and fibrosis

Autosomal recessive

Galactosemia (galactose-1phosphate uridyl transferase deficiency)

GALT

Hepatic failure with cirrhosis, renal failure, cataracts, cardiac failure

Ovarian failure with streak gonads

Autosomal recessive

Martsolf syndrome

RAB3GAP1

Short stature, microbrachycephaly, cataracts, abnormal facies with relative prognathism due to maxillary hypoplasia

“Primary hypogonadism”

Autosomal recessive

Nijmegen Syndrome

NBN

Chromosomal instability, immunodeficiency, hypersensity to ionizing radiation, malignancy

Ovarian failure (primary)

Autosomal recessive

Rothmund-Thompson syndrome

RECQL4

Skin abnormalities (telangiectasia, erythrema irregular pigmentation), short statue, cataracts, sparse hair, small hands and feet, mental retardation, osteosarcoma

Ovarian failure (primary hypogonadism or delayed puberty)

Autosomal recessive

Werner syndrome

WRN

Short stature, premature senility, skin changes (scleroderma)

Ovarian failure

Autosomal recessive

Autoimmune polyendocrinopathycandidiasis-extodermal dystrophy syndrome, APECED

AIRE

Candidiasis, Addison’s disease, hypoparathyroidism, type 1 diabetes, alopecia, vitiligo, ectodermal dystrophy, celiac disease and other intestinal dysfunctions, chronic atrophic, gastritis, chronic active hepatitis, autoimmune thyroid disorders, pernicious anemia

Ovarian failure

Autosomal recessive

Bloom syndrome

BLM

Chromosomal breakage leading to early onset of aging, short stature and elevated rates of most cancers

“Reduced fertility due to hypogonadism”

Autosomal recessive

Perrault syndrome, PS

HSD17B4, HARS2, CLPP, LARS2, C10orf2

Sensorineural deafness in both males and females, and neurological manifestations in some patients

Ovarian failure

Autosomal recessive

Central nervous system leukodystrophy and ovarian failure, ovarioleukodystrophy

EIF2B2, EIF2B4, EIF2B5

Neurological disorder characterized by involvement of the white matter of the central nervous system. When Leukodystrophies associated with premature ovarian failure referred to as ovarioleukodystroph

Ovarian failure

Autosomal recessive

amenorrhea, further proving impaired ability to prevent apoptosis.[56] DAZL is an autosomal homolog of the Y chromosome DAZ gene and is expressed limited to germ cells. Dazl+/ mice are subfertile and Dazl/ mice are infertile in both sexes. Tung et al. demonstrated that missense mutations P6H, N10C, I37A, R115G were associated with the age at menopause in women, of which N10C is responsible for sperm count in men [57]. Furthermore, Tung et al. [57]

proved that patients with heterozygous DAZL mutation were capable of having offspring, whereas those with homozygous mutations (i.e., females with p.R115G homozygous mutation and males with p.N10C mutation) experienced spontaneous POF at early age in female and showed no sperm in males. These phenotypes were consistent with those observed in mice. Genes expressed on somatic cells may also contribute to germ cell failure. Wilms tumor gene (WT1) locates to

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Sertoli cells in testes and granulosa cells in ovaries. Mice lacking Wt1 show normal germ-cell migration, but fail to develop urogenital ridge, predictably presenting with renal and gonadal agenesis. Mutation screening in WT1 conducted in sporadic POF females or, azoospermia/ severe oligozoospermic males revealed that variants in WT1 were related to germ cell failure.[58,59] Lack of Sf-1(encoded by NR5A1, as described before) in mice also led to complete gonadal agenesis. Mutations in NR5A1 were responsible for different phenotypes of 46,XY disorders of sex development (DSD) and 46,XX POF. Identical variants were shared among affected males and their female siblings.[60] In summary, perturbation of multiple genes could explain earlier repeats of germ cell failure in males and females in the same sibship.

Galactosemia Galactosemia is caused by deficiency of galactose I-phosphate uridyl transferase (GALT), encoded by a gene on 9p. Somatic features included renal, hepatic, and ocular defects. Kaufman et al. [61] reported POF in 12 of 18 galactosemic women, and Waggoner et al. [62] observed ovarian failure in 8 of 47 galactosemic women. Pathogenesis is presumed to involve galactose toxicity during infancy or childhood; elevated fetal levels of toxic metabolites. Consistent with this hypothesis, a neonate with galactosemia showed normal ovarian histology. Pathogenesis involves excess galactose toxicity impairing the folliculogenesis, inducing resistance to gonadotropins, and accelerating follicular atresia. Given the necessity for dietary treatment during childhood to prevent mental retardation, it seems highly unlikely that previously undiagnosed galactosemia would prove to be the cause of ovarian failure in women presenting solely with primary amenorrhea or POF. Of greater relevance to gynecology, therefore, was a report in 1989 by Cramer et al. [63] that GALT heterozygotes were at increased risk for POF; however, Kaufman et al. [64] failed to confirm the observation of Cramer et al. [63], and even Cramer et al. later failed to confirm perturbations of GALT abnormalities in a later study of women with early menopause (EM). Moreover, not all homozygotes for human galactosemia are abnormal, nor are transgenic mice in which GALT is inactivated.

Carbohydrate-Deficient Glycoprotein (Phosphomannomutase Deficiency) In type 1 carbohydrate-deficient glycoprotein (CDG) deficiency, mannose-6-phosphate cannot be converted to mannose-1-phosphate. These lipid-linked mannosecontaining oligosaccharides are also needed because secretory glycoproteins are lacking. Lacking these

glycoproteins, the phenotype is ovarian failure characterized by ovaries devoid of follicular activity. The molecular pathogenesis in the CDG gene located on 16p13 is usually a missense mutation. Neurologic abnormalities coexist, encompassing hypotonia, hyperreflexia, unprovoked eye movements, ataxia, joint contractions, epilepsy, and stroke-like episodes subcutaneous fat deposits, hepatomegaly, cardiomyopathy, pericardial effusion, and factor XI (clotting) deficiency.

Deficiency of 17a-Hydroxylase/17,20-Desmolase Deficiency (CYP17) Sex steroid synthesis requires intact adrenal and gonadal biosynthetic pathways. Various gene products (enzymes) are required to convert cholesterol to testosterone and androstenedione and, hence, estrogens. The various enzyme blocks in the adrenal/gonadal biosynthetic pathways have varying but predictable consequences, depending on their site in the biosynthetic pathway (see Fig. 1). The most common adrenal biosynthetic problem involves deficiency of 21- or 11b-hydroxylase, either of which causes pseudohermaphroditism. These disorders cause genital ambiguity because of virilization, but need not be considered in the differential diagnosis of XX gonadal dysgenesis. Perturbations of CYP17 and CYP19 (aromatase), however, may cause gonadal abnormalities without accompanying genital abnormalities. If the cytochrome P450 enzyme 17a-hydroxylase/ 17–20-lyase is deficient, pregnenolone cannot be converted to 17a-hydroxypregnenolone. If the enzyme defect were complete, cortisol, androstenedione, testosterone, and estrogens could not be synthesized; however, 11-deoxycorticosterone and corticosterone could. With compensatory increase in adrenocorticotropic hormone (ACTH secretion), 11-deoxycorticosterone and corticosterone increase to result in hypernatremia, hypokalemia, and hypervolemia. Hypertension occurs. Aldosterone is decreased, presumably because hypervolemia suppresses the renin-angiotensin system. Like most enzyme defects, 17a-hydroxylase deficiency is inherited in autosomal-recessive fashion. Females (46, XX) with 17a-hydroxylase deficiency have normal external genitalia, but at puberty they fail to undergo normal secondary sexual development (primary amenorrhea). Affected males (46,XY) usually have genital ambiguity (male pseudohermaphroditism) because partial expression of the gene can produce some androgens. Affected females are ordinarily encountered in the differential diagnosis of XX gonadal dysgenesis. Usually, hypertension is the major distinguishing feature. Oocytes appear incapable of spontaneously exceeding a diameter >2.5 mm, but ovaries nonetheless respond to exogenous gonadotropins.

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The CYP17 gene is located on 10q24–25, and its gene product is a cytochrome P450 enzyme. >20 different mutations have been identified, scattered among the 8 exons. Mutations include missense mutations, duplications, deletions, and premature protein truncation. Most mutations have been observed in only a single family, yet another example of molecular heterogeneity. An exception exists in Mennonites of Dutch origin, where a 4-base duplication in exon 8 accounts for most cases. This founder mutation originated in Friesland. A single gene (and enzyme) is responsible for both 17a-hydroxylase and 17,20-desmolase (lyase) actions (see Fig. 2). A few patients having deficiency of both 17ahydroxylase and 17,20 lyase activities have been analyzed, with mutations differing in those showing only deficient 17a-hydroxylase or showing both hydroxylase and desmolase defects. Site-directed mutagenesis in the rat gene indicates that mutations closer to the 5-end are more deleterious.

Aromatase Mutations (CYP 19) Conversion of androgens (Δ4-androstenedione) to estrogens (estrone) requires cytochrome P-450 aromatase, an enzyme product of a 40-kb gene located on chromosome 15q21.1. Deficiency of the aromatase enzyme in 46,XX individuals can be associated with clitoral hypertrophy or genital ambiguity, but 46,XX aromatase deficiency may be recognized as primary amenorrhea in otherwise normal females. Ito et al. [65] reported aromatase mutation (CYP19) in an 18-year-old 46,XX Japanese woman having primary amenorrhea and cystic ovaries. Compound heterozygosity

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existed for two point mutations in exon 10. The mutant protein had no activity. Conte et al. [66] reported aromatase deficiency in a 46,XX woman presenting with primary amenorrhea, elevated gonadotropins, and ovarian cysts. Compound heterozygosity also existed for two mutations in exon 10. One mutation was C1303T (cysteine rather than arginine); the other was G1310A (tyrosine rather than cysteine). Clitoral enlargement at puberty was reported by Mullis et al. [67]. No breast development occurred. Follicle-stimulating hormone (FSH) was elevated; estrone and estradiol were decreased. Multiple ovarian follicular cysts were evident. Compound heterozygosity existed in the CYP19 locus.

46,XX Agonadia In 46,XY agonadia, gonads are completely absent, not represented by a fibrous streak. Agonadia usually occurs in 46,XY individuals but, rarely, 46,XX cases exist. In agonadia (XY or XX) external genitalia are abnormal but female-like; no more than rudimentary Mullerian or wolffian derivatives are present. External genitalia usually consist of a phallus about the size of a clitoris, underdeveloped labia majora, and nearly complete fusion of labioscrotal folds. Thus external genitalia are usually female-like in appearance, albeit not normal. Somatic anomalies coexist in approximately one-half of cases: craniofacial anomalies, vertebral anomalies, and mental retardation. Sporadic 46,XX agonadia cases were reported by Duck and Levinson [68]. Mendonca et al. [69] reported agonadia without somatic anomalies in phenotypic sibs having

FIG. 2 Pivotal adrenal and gonadal biosynthetic pathways. Letters designate enzymes or activities required for the appropriate conversions. A, 20-Hydroxylase and 20,22-desmolase; B, 3β-ol-dehydrogenase; C, 17α-hydroxylase; D, 17,20-desmolase; E, 17-ketosteroid reductase; F, 21-hydroxylase; G, 11-hydroxylase; H, aromatase. In addition to these enzymes, steroid acute regulatory protein (StAR), designated I, is responsible for transporting cholesterol to the site of steroid biosynthesis. Finally, 17α-hydroxylase (C) and 17,20-desmolase (D) activities are actually governed by a single gene. Modified from Simpson JL, Elias S. Genetics in obstetrics and gynecology. 3rd ed. Philadelphia: W.B. Saunders Company; 2003.

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unlike chromosomal complements (46,XY and 46,XX). Kennerknecht et al. [70] reported agonadism, hypoplasia of the pulmonary artery and lung, and dextrocardia in discordant sex sibs (XX and XY).

Fragile X Syndrome and Expansion of Triplet Nucleotide Repeats (CGG) A molecular perturbation seemingly relevant to ovarian failure is expansion of triplet nucleotide repeats (CGG). The prototype is the fragile X syndrome, caused by mutation of the FMR1 gene on Xq27. “Fragile” refers to a tendency toward chromosomal breakage when affected cells are cultured in folic acid-deficient media. Various fragile sites exist in humans, but FRAXA and FRAXE are relevant to the present discussion. The molecular basis of FRAXA involves repetition of the triplet repeat CGG (CGG)n 200 times or more at a given site on the X long arm (Xq17) (Fig. 3). Ordinarily, the normal number of CGG repeats in males is only 6–50. Males with CGG expansion show mental retardation, characteristic facial features, and large testes. Heterozygous females having 50–200 repeats are said to have a premutation. During female (but not male) meiosis, the number of triplet repeats may increase (expand). Thus, a woman with a FRAXA premutation may have an affected son if the number of CGG repeats on the X, she transmits to her offspring, were to expand during meiosis to >200. These molecular changes are readily detectable by straightforward molecular studies. Females may also be affected (50% likelihood) if CGG expansion occurs, but the phenotype is less severe than in affected males. Females with the FRAXA premutation may show POF. Schwartz et al. [71] reported that fragile X carrier females (premutation) more often showed oligomenorrhea than

noncarrier female relatives (38% vs 6%). Murray et al. [72] analyzed 1268 controls, 50 familial POF cases, and 244 sporadic POF cases. Among familial cases, 16% showed a FRAXA premutation; among sporadic cases, only 1.6% showed POF. In the same sample, POF was not increased in FRAXE. (Nevertheless, FMR2 gene located in FRAXE is regarded to be associated with POF.) Its mutation frequency in women with POF was 1.5% compared to 0.04% in normal women [73]. An international collaborative survey [74] of 395 FRAYA premutation carriers revealed that 63 (16%) underwent menopause at younger than 40 years of age; only 0.4% of controls did. Consistent with these findings are observations that heterozygous FRAXA women respond poorly to ovulation-inducing agents, producing fewer oocytes and fewer embryos in assisted reproductive technologies (ARTs). Surprisingly, however, frequency of POF was not increased in 128 FRAXA cases having a full mutation. Additionally, it should be noted that the premutation frequencies differs in diverse ethnic groups. The prevalence of FMR1 premutation [12] is 3.3%–6.7% in Caucasian patients with POF, whereas it is 0.49% and 1.56% for Chinese [75] and Japanese POF patients, respectively. Although it is widely known that FRAXA is associated with POF, the underlying mechanism remains to be elucidated. Fragile X mental retardation protein (FMRP) is a RNA-binding protein encoded by FMR1 gene, responsible for transporting mRNA (including itself) out of the nucleus and repressing translation. People with FMR1 premutation shows elevated FMR1 mRNA levels but decreased FMRP level. Increased levels of expanded mRNA may produce functional toxicity, resulting in follicles impairment and atresia. [76] Certain deleterious genes might be functional at the premutation stage but suppressed when number of CGG repeats exceeds 200–230.

FIG. 3 Diagram of the FMR1 gene (also called FRAYA) and the first exon in normal, premutation, and full mutation alleles. The oval immediately to the left of the start site of transcription represents the promoter region of the FMR1 gene. The open symbol represents active transcription, and the black symbol, silenced transcription. The vertical lines indicate CGG trinucleotides upstream of the methionine codon (AUG) at the translocational start site. Reprinted with permission from Warren ST, Nelson DL. Advances in molecular analysis of fragile X syndrome. JAMA 1994;271(7):536–542.

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Myotonic Dystrophy and Expansion of Triplet Repeats (CTG) Myotonic dystrophy (MD) is an autosomal-dominant disorder characterized by muscle wasting (head, neck, extremities), frontal balding, cataracts, and male hypogonadism (80%) caused by testicular atrophy. The pathogenesis of MD involves nucleotide expansion of CTG in the 3¢ untranslated region of a gene located on chromosome 19. Normally, there are 5–27 CTG repeats. Heterozygotes usually have at least 50 repeats; severely affected individuals show 600 or more. Female hypogonadism is much less common than male hypogonadism. Despite frequent citations in texts, ovarian failure actually seems not to be well documented. Nonetheless, as in FRAXA, there is a poor response to ovulation induction regions. Sermon et al. [77] report fewer embryos per cycle than in standard ART; thus pregnancy rates in preimplantation genetic diagnosis are decreased.

Blepharophimosis-Ptosis-Epicanthus (FOXL2) Blepharophimosis-Ptosis-Epicanthus syndrome I (BPES-Type1) is an autosomal-dominant multiplemalformation syndrome long known to be associated with ovarian failure. POF is the usual clinical manifestation, rather than complete ovarian failure. In one unusual case, Fraser et al. [78] reported that the ovaries of an affected individual were unresponsive to gonadotropins. Sib-pair analysis using polymorphic DNA variants initially localized the gene to chromosome 3 (3q22 ! 24). Positional cloning revealed mutation in the winged helix/forkhead transcription factor gene (FOXL2), a truncated protein in each of four cases. Reported mutations include three cases with stop codons and one with 17-bp duplication causing a frameshift and, hence, the truncated gene product. This indicates that the mechanism of gene action is haploinsufficiency. To date, >100 unique intragenic FOXL2 mutations have been identified in patients with BPES, from different ethnic origin [79]. Among these mutations, 44% are frameshift mutations, 33% in-frame changes, 12% nonsense mutations, and 11% are missense mutations [79]. The FOXL2 gene is expressed in mesenchyme of mouse eyelids and in granulosa cells of adult ovarian follicles, consistent with the phenotype observed in perturbation of the human homolog BPE. The ovarian phenotype of FOXL2 mutations is variable. Some patients are diagnosed as POF with total absence of follicles, whereas others develop resistant ovary syndrome still with notable primordial follicles [78]. Arrest of follicular

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maturation or primordial to primary follicles was also observed [80,81]. Taken together, different mutations may exert different effect on FOXL2 protein, follicular defects and ovarian phenotypes. In addition, Mutations in FOXL2 can lead to POF without eyelid abnormalities although this seemingly accounts for only a small number of ostensibly nonsyndrome POF cases. De Baere et al. [82] tabulated FOXL2 mutations detected to date and offer genotype-phenotype correlations. The authors predict that a protein truncated before the poly-Ala tract will lead to POF.

Autosomal-Dominant POF Familial tendencies reminiscent of autosomaldominant inheritance have long been recognized in cytogenetically normal women who have POF but no other clinical abnormalities. Dominant tendencies are especially associated with multiple-system autoimmune abnormalities (ovarian, adrenal, thyroid); however, families with dominantly transmitted POF exist with no autoimmune problems. This is further distinct from familial transmission of terminal deletions of Xp or Xq. Thus, Coulam et al. [83] reported POF in sibs who had an affected mother and aunt. Affected individuals in more than one generation were reported by Starup and Sele and Austin et al. [84]. Dominant gene(s) causing POF may be more common than once believed. Testa and colleagues and Vegetti et al. [85] are systematically studying women with POF (menopause at younger than 40 years of age) who were ascertained in the northern Italian population. After excluding 10 cases with known etiologies (5 chromosomal, 3 prior ovarian surgery, 1 prior chemotherapy, 1 galactosemia), there remained 71 probands. Of the 71, 22 (31%) had other affected relatives. A later report by the authors encompassed 130 cases, 28.5% familial [85]. Patterns of inheritance observed were consistent with either autosomal or X-linked dominant inheritance; transmission through both maternal and paternal lineage was observed. Of 30 other women experiencing EM (40–50 years) but not POF per se, half showed other affected relatives. Thus POF and EM (40–50 years) may constitute the same phenomena. The two different phenotypes have been observed within a single kindred transmission through either a paternal or maternal relative. Subsequently, Bachelot et al. [86] performed a prospective and retrospective study involving 357 patients with POF, and found 14% with a positive family history. Most cases remain unexplained, suggesting our understanding about the genetic basis of POF is still incomplete.

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POLYGENIC FACTORS IN POF: NORMAL CONTINUATIONS VARIATION AS A CAUSE OF HERITABLE EM Oocyte number (reservoir) could be low in some women simply on statistical (stochastic) grounds. A normal distribution exists for essentially all anatomic traits (e.g., height) and physiologic (e.g., blood pressure) traits, and this principle surely applies to oocyte number and reservoir. Rodent strains show characteristic breeding duration, implying genetic control over either the rate of oocyte depletion or the number of oocytes initially present. That a normal distribution of germ-cell number exists in ostensibly normal females is difficult to prove in humans, but some ostensibly normal (menstruating) women can be expected to have decreased oocyte reservoir or increased oocyte attrition on a genetic basis. This genetic basis is presumably not a single gene but polygenic (see Simpson and Elias). Age at human menopause is heritable. Cramer et al. [87] took appropriate confounders (e.g., hysterectomy) into account in a case–control study of 10,606 US women. Women with EM (40–45 years) were age-matched with controls who were either still menstruating or who had experienced menopause after age 46. Of 129 EM cases, 37.5% had a similarly affected mother, sister, aunt, or grandmother. Only 9% of controls had such a relative [odds ratio after adjustment: 6.1, 95% confidence interval (CI): 3.9–9.4]. The odds ratio was greatest (9.1) for sisters and greater when menopause occurred before 40 years. Results of Cramer et al. [87] were confirmed by Torgerson et al. [88], who studied women undergoing menopause during the 5-year centile ages 45–49 years. The likelihood was increased that menopause would occur in a similar 5-year centile in their daughters. Twin studies also confirmed heritability of age at menopause. Snieder and colleagues [88a] studied 275 monozygotic (MZ) and 353 dizygotic (DZ) UK twin pairs. Correlation (r) for age at menopause was 0.58 for MZ and 0.39 DZ twins; heritability (h2) was calculated to be 63%. Treloar et al. [89] studied 466 MZ and 262 DZ Australian twin pairs. For age at menopause, correlation (r) was 0.49–0.57 for MZ and 0.31–0.33 for DZ twin pairs. Differences between MZ and DZ held when iatrogenic causes of menopause were taken into account. In recent years, genome-wide association studies were applied to explore the genetic components of POF, age at natural menopause, and EM. Multiple loci associated with age at menopause, DNA damage repair and replication, and immune function were identified [90]. Hypothesis that both EM and POF represent the left tail of menopause distribution, suggested that overlaps in polygenic etiology existed between EM and POF [91].

OVARIAN RESISTANCE Ovarian resistance implies that the genes necessary for ovarian differentiation are intact, but the ovary is unable to respond to nonovarian stimuli (e.g., FSH, LH). Only a few disorders of ovarian resistance are known. Many of the disorders discussed in the previous section could be the result not of ovarian failure per se but of ovarian resistance. The prototype example occurs in XX gonadal dysgenesis without somatic anomalies, one form of which proved to result from a mutation in the FSH receptor gene.

XX Gonadal Dysgenesis Due to FSHR Gene Mutation (C566T) In Finland, Aittomaki [81] and Aittomaki et al. [92,93] have identified this condition as the primary cause of XX gonadal dysgenesis. A nationwide search identified 75 patients fulfilling these diagnostic criteria: primary or secondary amenorrhea, serum FSH > 40 IU/mL. These 75 cases included 57 sporadic cases and 18 cases having one or more affected relative (7 different families). A preponderance of cases resided in north central Finland, a more sparsely populated part of the country. The frequency was 1 per 8300 liveborn Finnish females; the relatively high incidence was attributed to a founder effect. Segregation ratio of 0.23 for female sibs was consistent with autosomal-recessive inheritance, as was the high consanguinity rate (12%). In the initial analysis of Aittomaki [81] and Aittomaki et al. [92,93], a specific mutation was found in V566A of the FSHR gene, in six families. This cytosine-thymidine transition (C566T) was later found in additional families [92]. The C566T mutation has only rarely been detected in women with 46,XX ovarian failure who reside outside Finland. In the United States, Layman et al. [94] found no FSHR mutations in 35 46,XX women having hypergonadotropic hypogonadism (15 with primary amenorrhea, 20 with secondary amenorrhea). Liu et al. [95] found no FSHR anomalies in one multigenerational POF family, four sporadic POF cases, and two other hypergonadotropic hypogonadism cases. No cases were found in 46,XX POF or primary amenorrhea cases from Germany, Brazil, and Mexico. The last report analyzed all exons of FSHR. Similarly, Jiang et al. [96] failed to detect C566T in 1100 normal Danish or 540 normal Singaporean individuals (v. 1% in the general Finnish population; 1 in 1200 in Switzerland showed C566T heterozygosity). Comparing the phenotype of C566T XX gonadal dysgenesis (V566A) with non-V566A XX gonadal dysgenesis, Aittomaki et al. [92] showed the former to

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be more likely to have ovarian follicles on ultrasound examination. Thus C566T XX gonadal dysgenesis showed that some of the features gynecologists have long predicted for a gonadotropin-resistant disorder (Savage syndrome) although in general the phenotype of streak gonads found by Aittomaki [81] was not expected. FSHR (knock-out) mice similarly show failure of oogenesis; however, the necessity of FSH for progression of oogenesis is clear.

Inactivating Luteinizing Hormone Receptor Defect (46,XX) Another trophic hormone receptor gene whose perturbation causes XX gonadal dysgenesis is LHR, 75 kd in length and consisting of 17 exons. Located on 2p near FSHR, the first 10 exons of LHR are extracellular; the last 6 are intracellular; the 11th, transmembrane. LHR mutations have been reported predominantly in 46,XY individuals, where the phenotype may extend to complete LH resistance and XY gender reversal (female). 46,XX cases are less common than 46,XY cases, for reasons that are unclear. The only reported cases have occurred in sibships in which an affected 46,XY male had Leydig cell hypoplasia. 46,XX women with LHR mutations show oligomenorrhea or, less often, primary amenorrhea. Ovulation does not occur, although gametogenesis proceeds until the preovulatory stage but not beyond. This is consistent with findings in the mouse knockout model. Mutations in human LHR are molecularly heterogeneous, but most have been found in the transmembrane domain (exon 11). Latronico et al. [97] reported primary amenorrhea in a 22-year-old woman (46,XX) who had three affected male sibs (46,XY). Like her sibs, the 46,XX female was homozygous for a nonsense mutation at codon 554 (R554*). The resulting stop codon produced a truncated protein. The affected 46, XX female showed breast development, but only experienced a single episode of menstrual bleeding at age 20 years; LH was 37 mIU/mL, FSH 9 mIU/mL. The mutation reduced signal transduction activity of the LHR gene. Toledo and colleagues [98] studied a 46,XX female whose two 46,XY affected sibs had been previously reported by Cramer et al. [87] The sister showed elevated gonadotropins but anatomically normal ovaries. The mutation was A593P. Two sisters reported by Laue and colleagues showed the nonsense mutation C545* in exon 11; the father but not the mother had the mutation. The authors postulated a dominant negative effect, but more likely a mutant allele was also transmitted from the probably heterozygous mother but not recognized.

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[76] Mila M, Alvarez-Mora MI, Madrigal I, Rodriguez-Revenga L. Fragile X syndrome: an overview and update of the FMR1 gene. Clin Genet 2018;93(2):197–205. [77] Sermon K, De Vos A, Van de Velde H, Seneca S, Lissens W, Joris H, et al. Fluorescent PCR and automated fragment analysis for the clinical application of preimplantation genetic diagnosis of myotonic dystrophy (Steinert’s disease). Mol Hum Reprod 1998;4(8):791–6. [78] Fraser IS, Shearman RP, Smith A, Russell P. An association among blepharophimosis, resistant ovary syndrome, and true premature menopause. Fertil Steril 1988;50(5):747–51. [79] Verdin H, De Baere E. FOXL2 impairment in human disease. Horm Res Paediatr 2012;77(1):2–11. [80] Meduri G, Bachelot A, Duflos C, Bstandig B, Poirot C, Genestie C, et al. FOXL2 mutations lead to different ovarian phenotypes in BPES patients: case report. Hum Reprod 2010;25(1):235–43. [81] Aittomaki K. The genetics of XX gonadal dysgenesis. Am J Hum Genet 1994;54(5):844–51. [82] De Baere E, Beysen D, Oley C, Lorenz B, Cocquet J, De Sutter P, et al. FOXL2 and BPES: mutational hotspots, phenotypic variability, and revision of the genotype-phenotype correlation. Am J Hum Genet 2003;72(2):478–87. [83] Coulam CB, Stringfellow S, Hoefnagel D. Evidence for a genetic factor in the etiology of premature ovarian failure. Fertil Steril 1983;40(5):693–5. [84] Austin GE, Coulam CB, Ryan RJ. A search for antibodies to luteinizing hormone receptors in premature ovarian failure. Mayo Clin Proc 1979;54(6):394–400. [85] Vegetti W, Marozzi A, Manfredini E, Testa G, Alagna F, Nicolosi A, et al. Premature ovarian failure. Mol Cell Endocrinol 2000;161 (1–2):53–7. [86] Bachelot A, Rouxel A, Massin N, Dulon J, Courtillot C, Matuchansky C, et al. Phenotyping and genetic studies of 357 consecutive patients presenting with premature ovarian failure. Eur J Endocrinol 2009;161(1):179–87. [87] Cramer DW, Xu H, Harlow BL. Family history as a predictor of early menopause. Fertil Steril 1995;64(4):740–5. [88] Torgerson DJ, Thomas RE, Reid DM. Mothers and daughters menopausal ages: is there a link? Eur J Obstet Gynecol Reprod Biol 1997;74(1):63–6. [88a] Snieder H, MacGregor AJ, Spector TD. Genes control the cessation of a woman’s reproductive life: a twin study of hysterectomy and age at menopause. J Clin Endocrinol Metab 1998;83(6):1875–80. [89] Treloar SA, Martin NG, Heath AC. Longitudinal genetic analysis of menstrual flow, pain, and limitation in a sample of Australian twins. Behav Genet 1998;28(2):107–16. [90] Stolk L, Perry JR, Chasman DI, He C, Mangino M, Sulem P, et al. Meta-analyses identify 13 loci associated with age at menopause and highlight DNA repair and immune pathways. Nat Genet 2012;44(3):260–8. [91] Perry JR, Corre T, Esko T, Chasman DI, Fischer K, Franceschini N, et al. A genome-wide association study of early menopause and the combined impact of identified variants. Hum Mol Genet 2013; 22(7):1465–72. [92] Aittomaki K, Herva R, Stenman UH, Juntunen K, Ylostalo P, Hovatta O, et al. Clinical features of primary ovarian failure caused by a point mutation in the follicle-stimulating hormone receptor gene. J Clin Endocrinol Metab 1996;81(10):3722–6. [93] Aittomaki K, Lucena JL, Pakarinen P, Sistonen P, Tapanainen J, Gromoll J, et al. Mutation in the follicle-stimulating hormone receptor gene causes hereditary hypergonadotropic ovarian failure. Cell 1995;82(6):959–68. [94] Layman LC, Amde S, Cohen DP, Jin M, Xie J. The Finnish follicle-stimulating hormone receptor gene mutation is rare in north American women with 46,XX ovarian failure. Fertil Steril 1998;69(2):300–2.

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[95] Liu JY, Gromoll J, Cedars MI, La Barbera AR. Identification of allelic variants in the follicle-stimulating hormone receptor genes of females with or without hypergonadotropic amenorrhea. Fertil Steril 1998;70(2):326–31. [96] Jiang X, Morland SJ, Hitchcock A, Thomas EJ, Campbell IG. Allelotyping of endometriosis with adjacent ovarian carcinoma reveals evidence of a common lineage. Cancer Res 1998;58 (8):1707–12.

[97] Latronico AC, Anasti J, Arnhold IJ, Rapaport R, Mendonca BB, Bloise W, et al. Brief report: testicular and ovarian resistance to luteinizing hormone caused by inactivating mutations of the luteinizing hormone-receptor gene. N Engl J Med 1996;334(8):507–12. [98] Toledo SP, Brunner HG, Kraaij R, Post M, Dahia PL, Hayashida CY, et al. An inactivating mutation of the luteinizing hormone receptor causes amenorrhea in a 46, XX female. J Clin Endocrinol Metab 1996;81(11):3850–4.

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29 Environmental Contaminants and Ovarian Toxicity W.G. Foster, A.M. Gannon, H.C. Furlong INTRODUCTION The ovary is central for the production of gametes (oocytes) and the gonadal steroids estrogen and progesterone. The ovary is composed of a medulla surrounded by a cortex containing follicles at different stages of development. Follicles are composed of the theca externa and interna (androgen production), a basement membrane, granulosa cells (sites of estradiol (E2), and antim€ ullerian hormone (AMH) synthesis) and an oocyte. In mammals, oogenesis begins in utero with migration of the gametes from the germinal ridge into the primitive ovary. The first part of oogenesis starts in the germinal epithelium, which gives rise to the development of ovarian follicles, the functional unit of the ovary. Following puberty primordial follicles are recruited into the growing pool of follicles where they begin to grow through expansion of the granulosa cell population through a coordinated process that is initially gonadotropin independent. Follicle development is primarily driven by proliferation of granulosa cells becoming dependent on follicle-stimulating hormone (FSH) at the tertiary follicle stage shifting to luteinizing hormone (LH) at the time of ovulation and throughout the luteal phase of the menstrual cycle. Follicle recruitment, follicle development, ovulation, and luteal function are orchestrated through a complex system of feedback mechanisms of hormones, cytokines, and chemokines. Each step in the process can therefore potentially disrupted by exogenous chemicals; however, the extent to which environmental contaminants affect ovarian function remains poorly defined. There are more than 7 million known chemicals of which it is estimated that approximately 80,000 are in commercial use, rendering human chemical exposure inevitable. Human exposure to environmental contaminants has been widely documented and is widely

The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00029-7

thought to adversely affect human fertility. The global prevalence of infertility is estimated to be 9% [1] and between 50 and 80 million people worldwide are infertile [2]. Advanced age, diet, prescription medication use, preexisting health status, and infections are among the known risk factors for infertility; however, for many women the cause of their infertility is unknown. Environmental contaminants and life-style factors are thought to adversely affect human fertility in part via impaired ovarian function [3–5]. Among the many pathological and iatrogenic conditions that jeopardize reproductive function, chemical exposures have been associated with adverse health effects on the structure and function of the reproductive tract in females including: altered timing of pubertal onset, disruption of menstrual cycle function [6, 7], subfertility and impaired response to ovulation induction [8–10], ovarian cancer [11], increased time to achieve pregnancy [12], spontaneous abortion [13, 14], polycystic ovarian syndrome (PCOS) [15–17], endometriosis [18–21], and earlier age at menopause [22]. While the literature demonstrates numerous examples of potential associations between environmental contaminant exposure and adverse reproductive health outcomes, many others have been unable to show similar associations [23–25], hence links with adverse reproductive health outcomes in females remain uncertain. Although potential effects in the general population remain uncertain, the strongest evidence and potential for modifiable risk is limited to occupational groups (e.g., farmers) or specific populations whose lifestyle, such as sport fishing, presents risks for high exposure to known reproductive toxicants [26]. Therefore, in this chapter we review the biomonitoring, epidemiological, and experimental literature to explore the biological plausibility and elucidate potentially important mechanisms underlying potential associations.

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BIOMONITORING STUDIES AND EVIDENCE OF OVARIAN EXPOSURE Several studies have documented the concentration of environmental contaminants in ovarian follicular fluid of women undergoing fertility care. For example, benzene [9], persistent organic pollutants, including the polychlorinated biphenyl (PCB) congeners, polybrominated diphenyl ethers (PBDEs), and the pesticides such as: 1,1,1-trichloro-2,2-bis(p-chlorophenyl)ethane (DDT), the DDT metabolite 1,1-dichloro-2,2-bis(p-chlorophenyl)ethylene (DDE), and hexachlorobenzene, were quantified in the ovarian follicular fluid of women [10, 27–36]. Similarly, several other studies have also quantified the concentration of metals in ovarian follicular fluid [37–39]. Moreover, the concentration of polyaromatic hydrocarbons (PAHs) have also been measured in the follicular fluid of women who smoke tobacco vs. a control population undergoing assisted reproductive care [40]. Recent studies have also reported concentrations of chemicals with short half-lives that do not bioaccumulate in human tissues and are thought to act as endocrine toxicants. The concentrations of the perfluoroalkyl acids (PFAAs), such as perfluorooctanesulfonate, have been quantified in the ovarian follicular fluid of some women [41, 42]. Furthermore, the concentrations of Bisphenol A (BPA) and phthalates have also been measured in human follicular fluid of women attending assisted reproductive care centers [43, 44] although not all studies have been able to measure the concentrations of all compounds [45]. Collectively these studies establish that environmental contaminants are distributed to the ovary at concentrations high enough to be quantified in some women. Therefore, the consequences of environmental contaminant exposure on ovarian structure and function require further research.

EPIDEMIOLOGICAL EVIDENCE OF OVARIAN DYSFUNCTION Exposure to environmental contaminants is thought to pose a serious risk to human reproductive health. Exposure to a broad range of environmental contaminants including metals, pesticides, POPs, and nonpersistent endocrine toxic chemicals, has been linked to adverse reproductive outcomes that provide indirect evidence of potential effects on ovarian function. For example, increased time-to-pregnancy (TTP) has been detected in multiple studies investigating associations with life-style factors including cigarette smoking [46] as well as environmental contaminants including metals [47], PBDE’s [48], tetrachlorodibenzo[p]dioxin (TCDD) [49], PCB’s [50, 51], PFAA’s [52], BPA and phthalates [6, 53–55], parabens [56], and triclosan [54, 56]. Exposure to pesticides

has also been linked with decreased fecundity [12] whereas others have been unable to show a similar association although increased risk was demonstrated with older age in greenhouse workers [57]. Although adult exposure to pesticides was not linked with a delay in achieving pregnancy, daughters who were exposed in utero had an increased risk of delayed TTP [58]. In combination, these data suggest that age is a potential modifying factor or the effects of environmental contaminant exposure on TTP and raise the potential for transgenerational effects. Potential effects of environmental contaminants on ovarian function are also suggested by evidence of effects on menstrual cycle function. Disruption of menstrual cycle function has been associated with exposure to cigarette smoke [59], PCBs [7], DDT and DDE [7, 60], chlorinated by-products in drinking water [61], and BPA [6]. Further support for the hypothesis that environmental contaminants affect ovarian function comes from studies involving women receiving medical attention to achieve pregnancy. Specifically, in these studies exposure to BPA [62, 63] and phthalates [64] have been linked with a decrease in the number of eggs retrieved following controlled ovarian stimulation. Moreover, phthalate exposure has also been linked with decreased antral follicle counts [65] suggesting a decrease in follicle reserve. Ovarian toxicity of environmental contaminants is further advanced by studies demonstrating a potential link between exposure and changes in circulating estradiol [66], decreased fertilization rates [38], and oocyte quality [67]. While the epidemiological evidence suggests an association between exposure to environmental contaminants and impaired reproductive health, potentially through disruption of ovarian function, these studies have been criticized on several fronts. Specifically, these studies frequently measure multiple chemicals and examine the association between these chemicals and numerous reproductive outcomes. Thus, multiple hypotheses are often tested without correcting for multiple comparisons, increasing the risk of type I error (chance discovery). The strength of the associations found is generally weak and inconsistent across studies for individual compounds. Issues relating to exposure further complicate interpretation of study results. For example, age at exposure, dose, duration of exposure, and temporal relationship with the outcome measures are frequently unknown. While these studies endeavor to control for potential confounders this too represents an important challenge as comorbidities and underlying cause of infertility is commonly unknown. Consequently, translation of results from a population seeking medical intervention for infertility cannot easily be generalized to the broader population. Moreover, the concentrations measured in these studies are frequently in the pg/mL to low ng/mL range and

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association between these concentrations are many orders of magnitude below the concentrations needed to induce adverse effects in animal models. Finally, alternative explanations are frequently not considered. For example, phthalates are widely known to be present in medical devices including syringes, intravenous tubing, and bags. Thus, there is the potential that higher exposures in women who fail to conceive following medical intervention may be a marker of exposure to medical devices containing phthalates rather than an impact of the phthalates on ovarian function and fertility. Consequently, the relationship between exposure and adverse reproductive outcomes are viewed as limited.

BIOLOGICAL PLAUSIBILITY OF EFFECTS AND POTENTIAL MECHANISMS While the epidemiological literature provides limited support for the relationship between exposure to environmental contaminants and adverse effects on human fertility and ovarian function, animal studies provide support for the biological plausibility of the above associations. Experimental studies in animals reveal that environmental contaminants from all chemical classes, at some dose, can induce ovarian toxicity. Environmental contaminants inhibit germinal nest breakdown and induce oocyte aneuploidy [68, 69], produce anomalies of reproductive tract development [70–72], disrupt ovarian steroidogenesis [73], decrease ovarian weight or volume and ovarian follicle counts [74–76], and disrupt estrus (rodent) and menstrual (nonhuman primate) cycles. In a series of papers arising from the serendipitous observation of BPA-induced decrease in fertility, this nonpersistent contaminant induced adverse effects on germinal nest break down [68, 69, 77]. These results are important because they suggest that contaminant exposure can disrupt follicle formation leading to a decrease in the number of follicles in the resting pool. In mammals, females are born with a finite population of primordial follicles that represent the resting pool of follicles. In adult humans, a cohort of primordial follicles enters one of the waves of growing follicles with each menstrual cycle [78, 79]. Typically, one follicle from the growing pool of follicles is selected to ovulate while the remainder becomes atretic. Of the estimated 500,000 follicles present in the human ovary at the start of reproductive life, only about 400 reach the preovulatory stage and ovulate. Therefore, follicle atresia is the fate for the vast majority of female germ cells [80] and depletion of the resting pool either through disruption of the number of follicles formed, accelerated recruitment into the growing pool of follicles or increased destruction of follicles are all serious signs of ovarian toxicity.

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Folliculogenesis (follicle recruitment and growth to ovulation) is regulated in a stage-dependent manner by oocyte derived factors (e.g., bone morphogenetic protein-15 (BMP-15), growth differentiation factor-9 (GDF-9) and kit and kit ligand), gonadotropins (FSH and LH in early antral and Graafian follicles), and several growth regulators [e.g., transforming growth factor-β (TGF-β), AMH, inhibin-B, activin, vascular endothelial growth factor, FOXL2, and insulin-like growth factors and binding proteins], as previously reviewed in Ref. [81]. The mechanisms regulating primordial follicle recruitment are incompletely understood; however, recent studies demonstrate that AMH, a member of the TGF-β family, is produced exclusively by healthy growing follicles, is positively correlated with antral follicle count, and is a negative regulator of primordial follicle recruitment [82–86]. AMH is therefore thought to have clinical value as a marker of ovarian aging [82–90] and may also have value as a marker of ovarian toxicity. Of note, circulating levels of AMH decline with age [91] and are lower in women who smoke compared to nonsmokers [92, 93] although others have not found a similar relationship [94, 95], potentially owing to differences in methods and age of study subjects. Circulating concentrations of AMH were lower in mice exposed to cigarette smoke compared to controls [96] as well as in rat [97] and mouse [98] follicles cultured in the presence of B[a]P, a key chemical constituent of cigarette smoke. Methoxychlor treatment of postnatal day 4 rats, a stage during which primordial follicles are assembled, stimulated ovarian production of AMH [99], and parabens treatment induced an increase in AMH mRNA expression in neonatal rats [100]. While adverse effects of environmental contaminants are well established, the underlying mechanisms remain poorly defined. Mechanistic pathways inculpated in adverse effects of environmental contaminants included environmental contaminants or their metabolites binding with and activation of nuclear receptors including estrogen receptors (EsR1 and EsR2), androgen receptor (AR), and the AhR. Several environmental contaminants have estrogenic activity either through directly binding with the estrogen receptor or membrane receptors (GPR30), or estrogen-related receptor gamma (ERRγ). Several PCB congeners, pesticides, and nonpersistent contaminants, such as BPA, parabens, and triclosan bind with and activate estrogen receptors as shown by rodent uterotrophic and gene reporter assays. Phthalates and vinclozolin bind with the AR displacing testosterone, which is then aromatized to endogenous estrogens. TCDD, dioxin-like PCB congeners and PAHs bind with and activate the AhR, which has been linked with antiestrogenic effects. Contaminant binding with and activation of steroid receptors can disrupt ovarian follicle development, steroidogenesis, leading to oxidative stress, disruption

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of cell cycle kinetics, and triggering of cell death pathways including autophagy [101–103] and apoptosis [104–107]. Animal studies have also been key in elucidating the importance of developmental stage and suggesting the potential for adverse effects across multiple generations [108] through epigenetic modification of gene expression [109–113]. Consequently, animal studies have become indispensable in furnishing the scientific community with experimental evidence central to establishing biological plausibility. The experimental animal evidence elucidates dose response relationships that inform the assessment of human health risks and illustrate potential mechanisms of action. However, direct translation of results from animal studies to humans is challenging owing to differences in metabolic rate, metabolism of the test agents, physiology of the model, and underling mechanisms. For example, in general, some species such as mice are more sensitive than rats to chemicals acting through binding to and activation of the aryl hydrocarbon receptor [114]. Furthermore, the sensitivity to estrogens differs across species [115] and some strains of mice are more sensitive to estrogens than others [116]. Moreover, the mechanisms underlying the reproductive phenomena of interest may differ across test models, such as shown in Long Evans Hooded vs. Sprague-Dawley rats [117]. Hence, care must be assigned to selection of appropriate animal models for the goals of the study especially if mechanistic insight is needed. Accordingly, animal studies provide insight into potential health hazards and dose-response relationships are useful in assessing risk to health.

SUMMARY AND CONCLUSIONS Biomonitoring studies have established that environmental contaminants are distributed to the ovary and can be quantified in ovarian follicular fluid. Although epidemiological studies have shown associations between exposure to environmental contaminants and changes in circulating concentrations of gonadal steroids, increased TTP and infertility, other investigators have been unable to replicate these findings in similar study designs and populations and in some cases with higher exposures. Important limitations include the failure to account for multiple comparisons, confounding by co-exposure to other environmental contaminants, impact of comorbidities. Thus, the epidemiological studies provide only limited evidence for environmental contaminant-induced reproductive dysfunction and ovarian toxicity. However, the experimental animal evidence from in vivo and in vitro studies provides clear evidence for the biological plausibility of the associations

suggested in the epidemiological literature. Moreover, the experimental literature also elucidates potentially important mechanistic pathways. Moving forward it will be important to establish the effects of environmental contaminants at concentrations relevant to human exposure in the general population as well as those occupationally exposed. Use of innovative techniques, such as isolated ovarian follicle culture and organ on a chip, promise to provide greater mechanistic insight. We also suggest that attention should be given to investigating the effect of chemical interactions using mixtures of contaminants that reflect human exposure as recent studies suggest that chemical interactions may disrupt metabolism and increase target tissue concentrations [118, 119]. Contaminant-induced epigenetic modifications in the ovary is another area that holds great promise for aiding in understanding to chemical-induced dysregulation of ovarian function.

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C H A P T E R

30 Autologous Transplantation of Human Ovarian Tissue Enes Taylan, Kutluk Oktay INTRODUCTION Throughout its physiological lifetime from intrauterine period to menopause, human ovary undergoes a unique developmental journey. Approximately at the 7th week of embryological development, the primitive ovary begins to originate from coelomic epithelium that lines the body cavity in the ventromedial surface of the mesonephros. During this transformative process, germ cells begin to migrate to the specified gonad. Mitotically active oogonial cells rapidly proliferate and colonize the ovary reaching 6–7 million in numbers at 20 weeks of gestation. From this point on, the number of oogonial cells gradually decreases due to significant follicle atresia. This results in only about 1 million germ cells remaining at birth. Of those only 300,000–500,000 primordial follicles remain at pubertal age [1]. Following puberty, this number is gradually reduced to about 25,000 by the age 37 years followed by an accelerated phase of loss and nearly exhausted at menopause. While the inefficiency of the maintenance of ovarian reserve may not bode well for the reproductive lifespan, the redundancy in primordial follicle numbers lends itself to ovarian tissue cryopreservation technologies. In the last two decades, survival rates among children and reproductive age women with cancer have significantly improved, largely due to the development of more effective cancer diagnosis and treatments. In addition to malignancies, a number of nononcological systemic diseases such as autoimmune and hematological disorders may also require chemotherapy or radiotherapy as conditioning treatments prior to bone marrow transplantation [2]. However, cancer a great majority of chemotherapy treatments and radiotherapy have irreversible effects on ovarian reserve [3–5]. Exposure to pelvic radiotherapy and chemotherapeutic agents, especially of the alkylating family induce DNA damage and apoptosis in primordial follicles [6], causing reduction and premature exhaustion

The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00030-3

of ovarian reserve [7,8]. Moreover, in some genetic disorders such as in Turner Syndrome, there is an accelerated follicular atresia that results in markedly reduced ovarian reserve at birth [9,10]. While oocyte or embryo cryopreservation is considered not experimental, their utility is limited in the face of imminent chemotherapy and/or radiotherapy and in children. Ovarian cryopreservation has evolved to fill that gap to preserve fertility in young women and children facing fertility damaging treatments. Ovarian tissue cryopreservation enables preservation of a large number of primordial follicles embedded in ovarian cortex before such gonadotoxic treatments. This procedure can be done at anytime of the menstrual cycle without need for ovarian stimulation. It typically requires surgically harvesting of ovarian tissue via a simple laparoscopic outpatient procedure followed by separation of stromal tissue from ovarian cortex that comprises primordial follicles. The cortical tissue is then cut into small pieces of 5  5  1 mm (width  length  thickness) and processed with various cryoprotectants. The currently established technique of freezing is Slow Freezing, but strides are being made with vitrification as well [11,12]. When the patient is cured of her disease and desires fertility, and in some cases for restoration of endocrine function, the tissues are thawed and autotransplanted. In this chapter, we will review the development and techniques of the ovarian autotransplantation procedure, its current state, and future aspects.

A BRIEF HISTORY OF OVARIAN TISSUE CRYOPRESERVATION AND TRANSPLANTATION For reproductive age women undergoing gonadotoxic therapies, there are several fertility preservation strategies. Oocyte and embryo cryopreservation are endorsed as established options by the American Society for

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Reproductive Medicine (ASRM) and the American Society of Clinical Oncology (ASCO) [13,14]. However, for women who cannot delay their treatment and prepubertal girls who cannot undergo oocyte or embryo cryopreservation, the only available option is ovarian tissue cryopreservation. Major indications for ovarian tissue cryopreservation and transplantation are summarized in Table 1. The first animal studies of ovarian tissue cryopreservation were performed by Parkes using slow freezing procedure in a rodent model in the early 1950s [15]. This procedure was improved and taken several steps further in the next 50 years by animal studies and human ovarian tissue xenograft models [16–18]. In 2000, Oktay et al. reported the first successful transplantation of frozenthawed human ovarian tissue, which showed restoration of endocrine function with follicular development in response to gonadotropin stimulation [19]. In their report, 80 pieces of ovarian tissue were thawed, sutured to two triangular frames made from an absorbable cellulose membrane (Surgicel, Ethicon, Somerville, NJ) and laparoscopically transplanted beneath the left pelvic sidewall peritoneum. Approximately 4 months after the transplantation, daily gonadotropin administration was started, and after 24 days of stimulation, a follicle reached 17 mm in diameter with appropriate estradiol production. Ovulation was triggered by human chorionic gonadotropin (h-CG) administration and confirmed by

ultrasound imaging with subsequent rise and peak in serum progesterone. Follicular development was still observable in the graft via ultrasonography 6 months after the transplantation. Following this achievement, Oktay et al. reported oocyte retrieval and embryo development after heterotopic transplantation of ovarian tissue under forearm and abdominal skin [20,21]. In the ensuing year, over 80 livebirths have been reported using this and the variations of the ovarian autotransplantation techniques [22].

TRANSPLANTATION TECHNIQUES OF FROZEN-THAWED OVARIAN TISSUE Once the antineoplastic treatment is completed and the patient is cured of her disease, the previously frozen ovarian tissue can be thawed and transplanted via orthotopic or heterotopic transplantation methods [22] (Table 2). TABLE 2

Comparison of Ovarian Tissue Transplantation Techniques Orthotopic Ovarian Transplantation

Heterotopic Ovarian Transplantation

Reported transplantation sites

• In situ ovary • Beneath the pelvic sidewall peritoneum • Beneath the fallopian tube serosa

• Subcutaneous (lower abdomen/forearm) • Subperitoneal, beneath rectus muscle • Between breast tissue and pectoralis muscle

Pros

• Physiological microenvironment for follicular development • Possibility of natural conception

• No need for entry into abdominal/pelvic cavity, less invasive • No need for general anesthesia • Can be done in office setting; lower cost • Ease of monitoring the graft in case of recurrence concern

Cons

• Requires entry into pelvic/abdominal cavity • General anesthesia is required • Potential for direct seeding of residual cancer cells into abdominal/pelvic cavity • Higher cost/need for hospital facilities

• IVF is required for conception • Microenvironment may be less favorable for follicular development

Current success rate

• Cumulative livebirth and ongoing pregnancy rate of 37.7%

• Unknown, as there are too few reports

TABLE 1 Examples of Cancer and Noncancer Indications for Ovarian Tissue Cryopreservation and Transplantation. Malignancies

Nonmalignant Conditions

Malignancies common in young females • Leukemia • Hodgkin and non-Hodgkin lymphoma • Bone tumors (osteosarcoma, Ewing sarcoma) • Rhabdomyosarcoma • Wilms tumor • Neuroblastoma

Autoimmune and hematological diseases that require chemotherapy or HSCT • Systemic lupus erythematosus • Rheumatoid arthritis • Acute glomerulonephritis • Behc¸et’s disease • Aplastic anemia • Sickle cell anemia • Severe Combined Immunodeficiency Syndrome

Breast cancer (Stage I–-III) Gynecological cancers • Early stage cervical cancer • Endometrial cancer (no ovarian involvement) • Ovarian cancer (borderline tumors, early stage) Diseases that require pelvic radiation • Pelvic solid organ tumors (gynecologic, gastrointestinal, or genitourinary malignancies) • Pelvic bone tumors • Tumors of the spinal cord

Benign ovarian diseases • Endometriosis • Benign ovarian tumors Prophylactic oophorectomy • BRCA 1 and 2 germline mutation carriers • P53 mutation carriers • Carriers of other gynecologic cancer predisposing gene mutations Genetic disorders that cause premature ovarian failure • Mosaic Turner Syndrome • FMR1 premutation

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TRANSPLANTATION TECHNIQUES OF FROZEN-THAWED OVARIAN TISSUE

Primordial follicle embedded in the ovarian cortex

Fallopian tube

B Mesosalpinx Uterus A Ovary

Tissue grasper

C

D Reconstructed ovarian graft sutured onto the bivalved ovary

Pelvic side wall peritoneum

FIG. 1 Orthotopic ovarian tissue transplantation techniques. (A) Transplantation under the cortex of the remaining ovary; (B) between the layers of mesosalpinx; (C) under the pelvic side wall peritoneum; (D) on to the bivalved remaining ovary.

In general, orthotopic ovarian transplantation refers to surgical techniques that can enable spontaneous conception and hence pelvic in location (Fig. 1). Heterotopic transplantation refers to other locations, usually outside the pelvis and sometimes in the upper extremity (Figs. 2 and 3). The primary choice for ovarian transplantation is orthotopic (pelvic) as this technique most closely mimics the natural function of the ovary and potentially allows spontaneous conception. Heterotopic ovarian transplantation is considered when pelvis is not suitable due to prior radiotherapy or scarring, existing morbidities make major pelvic surgery contraindicated, when there is a risk of primary disease recurrence in the frozen ovarian tissue after transplantation and patient choice, whether only endocrine function is desired [23]. Hence, ovarian transplantation technique is tailored based on individual circumstances of each case. A fundamental variable that influences the success of the transplantation and strongly determines the graft lifespan is the primordial follicle density in thawed ovarian

cortical tissue. In our practice, we always evaluate the primordial follicle density (number of primordial follicles per mm3 cortical tissue) in a single piece of frozen-thawed ovarian tissue before attempting ovarian transplantation. Not only this evaluation affirms the presence of primordial follicles prior to transplantation, but primordial follicle density guides as to how much tissue to thaw. As important is the preoperative histological evaluation of cryopreserved tissue to screen for residual tumor cells in patients with cancer history.

Orthotopic Ovarian Transplantation Orthotopic transplantation of ovarian tissue is defined as the surgical procedure in which the ovarian graft is transplanted back to the pelvic cavity where ovaries naturally reside. In this approach, ovarian cortical tissue is engrafted into the anatomical locations such as the remaining ovary, pelvic sidewall, and fallopian tube serosa. Because ovarian transplantation is performed

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30. AUTOLOGOUS TRANSPLANTATION OF HUMAN OVARIAN TISSUE

Rectus abdominus muscle Laparoscopic grasper Abdominal wall peritoneum Ovarian graft Internal oblique muscle Inferior epigastric artery

Pelvic cavity

Bladder

Rectum

Uterus

FIG. 2 Heterotopic transplantation of ovarian tissue to abdominal wall retroperitoneum.

Tissue grasper Ovarian graft Forearm skin

FIG. 3 Heterotopic transplantation of ovarian tissue under forearm skin.

without vascular anastomosis, reimplantation on a vascular bed to facilitate tissue revascularization is crucial for the survival of the graft. These anatomical sites possess vascular networks that can provide sufficient oxygen and nutrient supply [24]. Ovarian cortical tissues can be cryopreserved in size ranging from 2 to 3 mm2 to 1 cm2, though we prefer 0.25 mm2 pieces (5  5 mm pieces in 1 mm thickness) as this lends itself to easier manipulation and assembly prior to transplantation [19,25].

In the case when there is at least one remaining ovary with intact arterial supply, cryopreserved ovarian tissues can be successfully transplanted to this recipient ovary using a number of methods. In one approach, after removal of a large piece of cortex from recipient ovary, frozenthawed ovarian tissues are fixed to the exposed ovarian medulla using 7-0 or 8-0 propylene sutures. In another technique, ovarian cortical pieces are directly deposited in the medulla after creating an incisional hole on the recipient ovary cortex [26]. However, a limited amount of ovarian tissue can be transplanted in both techniques. To maximize the surface area on the recipient ovary, we have developed a new technique that is performed by bivalving the ovary [27]. In our technique, bivalving of the recipient ovary exposes the vascular medulla and provides a larger surface area for the transplanted graft [28,29]. When there is no remaining ovary or the remaining ovary appears atrophic or nonfunctional, ovarian cortical pieces can be grafted the pelvic sidewall peritoneum [19] or the fallopian tube [25]. Since the first successful surgery performed by Oktay et al. in 2000, orthotopic ovarian transplantation has been the main approach, as a result, being associated with

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LIMITATIONS WITH OVARIAN TRANSPLANTATION

nearly all livebirths after this procedure [19,27,30]. The success rates of ovarian autotransplantation will be discussed later in this chapter.

Heterotopic Ovarian Transplantation In certain instances, orthotopic ovarian transplantation may not be feasible, necessitating a heterotopic approach (Table 2). In the heterotopic approach, the tissues are transplanted outside the pelvic cavity. These sites include forearm or abdominal subcutaneous locations, rectus fascia, and retroperitoneally in the upper abdomen [20,21]. These heterotopic sites have been shown to provide viable microenvironment for graft survival under sufficient vascular supply [31]. In humans, we have previously reported the restoration of endocrine function, follicular development, and embryo generation following gonadotropin stimulation after subcutaneous transplantation of ovarian tissues [20,21]. We showed that ovarian tissue can be transplanted subcutaneously, under local anesthesia only. Because of its ease and simplicity, a heterotopic ovarian transplantation can be performed repetitively as needed on the same patients and may be more cost-effective compared to orthotopic transplantation. However, the heterotopic sites may not provide the physiological paracrine milieu and temperature for optimal follicle development. For example, in our early experiences, follicles that were transplanted under the skin only grew to 10–14 mm sizes before yielding mature oocytes and embryo yield was very low. While we cannot rule out that this was due to the quality of freezing, there is strong suspicion that superficial ovarian transplantation may affect oocyte quality. However, intraabdominalretroperitoneal heterotopic transplant techniques may overcome those shortcomings. Hence, for patients whose primary goal is to restore fertility, orthotopic ovarian transplantation is preferred, unless there is a medical contraindication. If the main goal is to restore, ovarian endocrine function, heterotopic technique may be more practical. Although spontaneous pregnancy is not expected after heterotopic ovarian transplantation, we reported a case with multiple spontaneous conceptions and livebirths after heterotopic transplantation of ovarian tissue beneath the suprapubic skin [32,33]. After subcutaneous transplantation, the patient’s graft began functioning and resulted in follicle development as well as oocyte retrieval. While this oocyte was not fertilized, the patient spontaneously conceived in the subsequent cycle but had a miscarriage. She then had 3 consecutive spontaneous pregnancies, which all resulted in livebirth. Given that she had had bone marrow transplantation and had been menopausal for 2.5 years, it was highly implausible that

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this was a spontaneous event. Further research needs to explore, whether transplanted ovary can have a regenerative effect on the remaining menopausal ovary. Recently, Stern et al. reported the first livebirth following IVF after heterotopic transplantation of frozen-thawed ovarian tissues retroperitoneally beneath the abdominal wall [34]. Although this is the only published livebirth after a heterotopic ovarian transplantation procedure, others have been reported in abstract presentations [35], and more progress is underway with this technique. Depending on the initial follicle reserve of the tissue, it has been shown that the heterotopic graft may function up to 7 years [36]. Hence, further clinical studies with longer follow-up durations are needed to assess the efficiency and success of this technique in comparison to orthotopic transplantation method. Table 2 summarizes the pros and cons of heterotopic ovarian transplantation in comparison to orthotopic ovarian transplantation.

CURRENT SUCCESS RATE OF OVARIAN TRANSPLANTATION Cryopreservation and autotransplantation of ovarian tissue is still considered as an experimental procedure for fertility preservation. This is mainly due to the lack of systematic data that provides the actual success rate of this promising reproductive technology. Although remarkable progress has been witnessed in the last 20years, most of the published studies are case reports or case series that were not capable of clearly demonstrating the efficiency of ovarian transplantation. In a very recent meta-analysis, we were able to determine the current success rate of ovarian tissue transplantation with previously cryopreserved tissue [30]. After a comprehensive literature review from 1999 to 2016, we identified 309 ovarian tissue transplantation cases with cryopreserved ovarian tissue, which resulted in 84 livebirths and 8 ongoing pregnancies. The endocrine function restoration, cumulative clinical pregnancy, and livebirth plus ongoing pregnancy rates were 63.9%, 57.5%, and 37.7%, respectively. Furthermore, over 60% of those who underwent orthotopic ovarian transplantation could conceive naturally. These results clearly demonstrate the efficiency of ovarian transplantation and encourage us to consider this technology among the other established fertility preservation options.

LIMITATIONS WITH OVARIAN TRANSPLANTATION Ovarian cortical pieces are transplanted with a method similar to skin grafting, without vascular anastomosis. Because full revascularization and maturation of the new blood vessels take up to 10 days, nearly two-third

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of the ovarian reserve may be lost during this initial ischemic phase [37]. As a result, the longevity of ovarian transplants is generally limited, on average graft function ceasing in 26.9 months [30,38]. In our laboratory, we have been working on improving this revascularization process to enhance outcomes within the past decade. For example, we have shown that a ceramide-induced death inhibitor sphingosine-1phosphate (S1P) can accelerate ovarian graft revascularization and reduce ovarian ischemia while reducing primordial follicle loss when continuously infused during the first 3 days after the transplant in human ovarian xenograft models [39]. However, S1P has a very short half-life, not approved for human use and would need to be continuously infused during the first 3–4 days after the ovarian transplantation in humans, which makes it not practical. FTY 720 is a synthetic analogue of S1P, which is currently approved for medical treatment of multiple sclerosis. While our original studies did not find the same benefit in ovarian revascularization, further dose adjustment studies may result in the clinical use of this drug to enhance ovarian transplantation success. Another potential source of improvement may be through extracellular matrix scaffold. We have previously shown that the extracellular matrix plays an important role in primordial follicle growth and survival [40,41]. Using these data from earlier animal studies, we began utilizing an extracellular matrix scaffold to reconstruct previously frozen ovarian tissue prior to ovarian transplant. We combined this approach with robotic surgery to increase surgical precision and speed [11,27]. Initial surgical cases resulted in robust ovarian function and livebirths. As the number of transplant cases increases, we will be able to determine whether use of extracellular matrix scaffolds and robotic surgery improve ovarian transplant success and longevity. Vascular endothelial growth factor (VEGF) has been shown in some animal models to facilitate graft revascularization [42]. Among the other proposes pharmacological interventions are gonadotropins, fibroblast growth factor 2 (FGF2), angiopoietin-2, simvastatin, vitamin E, melatonin, and N-acetylcysteine. These have been shown to induce neovascularization and decrease ischemic graft necrosis in animal experiments [43–49]. However, none of these have been translated to the clinical field. With the application of current techniques, the mean duration of graft function after orthotopic ovarian transplantation appears to be 26.9 months (range: 4–144 months) based on available data [30].

RISK OF CANCER CELL REIMPLANTATION There has been a general concern with the risk of introducing cancer cells back into body along with

reimplantation of cryopreserved ovarian tissue. However, studies showed no evidence of malignant cells in cryopreserved ovarian tissues from patients with nonmetastatic solid tumors such as breast cancer and those with bone and soft tissue tumors [50–52]. One of the most concerning malignancies from the point of reseeding cancer cells along with ovarian tissue is acute leukemia. In patients with acute leukemia, the malignant cells present in circulation and hence in ovarian vessels as well. However, the chemotherapy regimens used in acute leukemia treatment are generally not gonadotoxic, and hence fertility preservation is not needed at that stage. Fertility preservation by ovarian tissue cryopreservation is needed when they are in remission and about to undergo hematopoietic stem cell transplantation (HSCT). The preconditioning chemotherapy prior to HSCT nearly guarantees ovarian failure. In such patients with remission, there may be very few cells present in circulation if any, and hence the risk of reseeding cancer via autotransplantation of frozen-thawed tissue is likely small. In fact, a human ovarian xenograft study found no risk of transmission from such patients by real-time PCR analysis of leukemia cells in thawed tissues and observation of cancer transmission to immunodeficient mice [53]. Regardless of the probability of ovarian involvement, a sample of thawed ovarian tissue should be histologically evaluated, and when available screened with molecular markers of the cancer of interest [54].

CONCLUSION AND FUTURE DIRECTIONS Ovarian tissue cryopreservation and transplantation is a fertility preservation approach that offers a lot more than the gamete freezing techniques. It can restore natural fertility and “reverse” menopause. Since the first successful report by our group, we have witnessed remarkable progress in cryopreservation methods and transplantation techniques. Although it is still defined as an experimental procedure, ovarian transplantation technology is rapidly evolving and the current success rates may be comparable to oocyte freezing. As a result, several countries outside of United States have taken ovarian tissue freezing from the experimental category. With further improvements in ovarian revascularization, ovarian autotransplantation may even further excel in efficiency. Though it is not currently possible to perform whole ovary transplantation in human, if that becomes possible 1 day, the ischemic losses may be minimized [55]. Research is also progressing with the vitrification of ovarian tissue [56,57]. If vitrification is successful, this may make ovarian freezing more practical and utilized more widespread. Other areas of research include screening and isolation of cancer cells in ovarian tissue prior to ovarian transplantation. If realized, all of these

V. HUMAN OVARIAN PATHOPHYSIOLOGY: SELECT ASPECTS

REFERENCES

areas of development may make ovarian tissue freezing and transplantation an effective method of fertility preservation not only for cancer patients but also for elective reasons as well.

Glossary Chemotherapy a type of cancer treatment in which cytotoxic drugs are used systemically. Cryopreservation a specific procedure in which various chemical solutions and antifreezing molecules are used to preserve cells or tissues by cooling to very low temperatures below zero. Gonadotoxic potentially damaging to gonads such as ovary and testis. Ovarian germ cell an ovarian cell with half number of chromosomes of somatic cell that unites with sperm cell to form a new individual. Primordial follicle an ovarian follicle consists of an oocyte surrounded by a single layer of granulosa cells. Radiotherapy a type of cancer treatment in which X-rays are used to treat abnormal growths.

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[13] Loren AW, Mangu PB, Beck LN, Brennan L, Magdalinski AJ, Partridge AH, Quinn G, Wallace WH, Oktay K. American society of clinical oncology. fertility preservation for patients with cancer: American Society of Clinical Oncology clinical practice guideline update. J Clin Oncol 2013;31:2500–10. [14] Ethics Committee of American Society for Reproductive Medicine. Fertility preservation and reproduction in patients facing gonadotoxic therapies: a committee opinion. Fertil Steril 2013;100:1224–31. [15] Parkes AS, Smith AU. Regeneration of rat ovarian tissue grafted after exposure to low temperatures. Proc R Soc Lond B Biol Sci 1953;140(901):455–70. [16] Deanesly R. Immature rat ovaries grafted after freezing and thawing. J Endocrinol 1954;11(2):197–200. [17] Newton H, Aubard Y, Rutherford A, Sharma V, Gosden R. Low temperature storage and grafting of human ovarian tissue. Hum Reprod 1996;11(7):1487–91. [18] Oktay K, Newton H, Mullan J, Gosden RG. Development of human primordial follicles to antral stages in SCID/hpg mice stimulated with follicle stimulating hormone. Hum Reprod 1998;13(5):1133–8. [19] Oktay K, Karlikaya G. Ovarian function after transplantation of frozen, banked autologous ovarian tissue. N Engl J Med 2000;342:1919. [20] Oktay K, Economos K, Kan M, Rucinski J, Veeck L, Rosenwaks Z. Endocrine function and oocyte retrieval after autologous transplantation of ovarian cortical strips to the forearm. JAMA 2001;286(12):1490–3. [21] Oktay K, Buyuk E, Veeck L, Zaninovic N, Xu K, Takeuchi T, Opsahl M, Rosenwaks Z. Embryo development after heterotopic transplantation of cryopreserved ovarian tissue. Lancet 2004;363(9412):837–40. [22] Akar M, Oktay K. Restoration of ovarian endocrine function by ovarian transplantation. Trends Endocrinol Metab 2005;16: 374–80. [23] Sonmezer M, Oktay K. Orthotopic and heterotopic ovarian tissue transplantation. Best Pract Res Clin Obstet Gynaecol 2010;24(1):113–26. [24] Oktay K, Buyuk E. Ovarian transplantation in humans: indications, techniques and the risk of reseeding cancer. Eur J Obstet Gynecol Reprod Biol 2004;113:S45–7. [25] Suzuki N, Yoshioka N, Takae S, Sugishita Y, Tamura M, Hashimoto S, Morimoto Y, Kawamura K. Successful fertility preservation following ovarian tissue vitrification in patients with primary ovarian insufficiency. Hum Reprod 2015;30(3): 608–15. [26] Andersen CY, Rosendahl M, Byskov AG, Loft A, Ottosen C, Dueholm M, Schmidt KL, Andersen AN, Ernst E. Two successful pregnancies following autotransplantation of frozen/thawed ovarian tissue. Hum Reprod 2008;23(10):2266–72. [27] Oktay K, Bedoschi G, Pacheco F, Turan V, Emirdar V. First pregnancies, live birth, and in vitro fertilization outcomes after transplantation of frozen-banked ovarian tissue with a human extracellular matrix scaffold using robot-assisted minimally invasive surgery. Am J Obstet Gynecol 2016;214: 94:e1–9. [28] Taylan E, Oktay KH. Robotics in reproduction, fertility preservation, and ovarian transplantation. Robot Surg Res Rev 2017;4:19–24. [29] Oktay K, Taylan E, Sugishita Y, Goldberg GM. Robot-assisted laparoscopic transplantation of frozen-thawed ovarian tissue. J Minim Invasive Gynecol 2017;24:897–8. [30] Pacheco F, Oktay K. Current success and efficiency of autologous ovarian transplantation: a meta-analysis. Reprod Sci 2017;24 (8):1111–20. [31] Suzuki N, Hashimoto S, Igarashi S, Takae S, Yamanaka M, Yamochi T, et al. Assessment of long-term function of heterotopic transplants of vitrified ovarian tissue in cynomolgus monkeys. Hum Reprod 2012;27:2420–9.

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[32] Oktay K. Spontaneous conceptions and live birth after heterotopic ovarian transplantation: is there a germline stem cell connection? Hum Reprod 2006;21(6):1345–8. [33] Oktay K, T€ urkc¸€ uo glu I, Rodriguez-Wallberg KA. Four spontaneous pregnancies and three live births following subcutaneous transplantation of frozen banked ovarian tissue: what is the explanation? Fertil Steril 2011;95(2) 804.e7–10. [34] Stern CJ, Gook D, Hale LG, Agresta F, Oldham J, Rozen G, Jobling T. Delivery of twins following heterotopic grafting of frozen-thawed ovarian tissue. Hum Reprod 2014;29(8):1828. [35] Stern K, Rozen G, Gook D, Agresta F, Braat D, Hale L. Are there factors which predict the success of ovarian tissue grafting in oncofertility patients? Abstract presented at: 32nd annual meeting of the European Society of Human Reproduction and Embryology; 2016. Helsinki, Finland. [36] Kim SS. Assessment of long-term endocrine function after transplantation of frozen-thawed human ovarian tissue to the heterotopic site: 10-year longitudinal follow-up study. J Assist Reprod Genet 2012;29(6):489–93. [37] Gosden RG, Baird DT, Wade JC, Webb R. Restoration of fertility to oophorectomized sheep by ovarian autografts stored at -196 degrees C. Hum Reprod 1994;9:597–603. [38] Lee J, Kong HS, Kim EJ, et al. Ovarian injury during cryopreservation and transplantation in mice: a comparative study between cryoinjury and ischemic injury. Hum Reprod 2016;31(8):1827–37. [39] Soleimani R, Heytens E, Oktay K. Enhancement of neoangiogenesis and follicle survival by sphingosine-1-phosphate in human ovarian tissue xenotransplants. PLoS One 2011;29: 6-e19475. [40] Oktay K, Karlikaya G, Akman O, Ojakian GK, Oktay M. Interaction of extracellular matrix and activin-A in the initiation of follicle growth in the mouse ovary. Biol Reprod 2000;63:457–61. [41] Oktem O, Oktay K. The role of extracellular matrix and activin-A in in vitro growth and survival of murine preantral follicles. Reprod Sci 2007;14:358–66. [42] Wang L, Ying YF, Ouyang YL, Wang JF, Xu J. VEGF and bFGF increase survival of xenografted human ovarian tissue in an experimental rabbit model. J Assist Reprod Genet 2013;30(10):1301–11. [43] Gao J, Huang Y, Li M, Zhao H, Zhao Y, Li R, Yan J, Yu Y, Qiao J. Effect of local basic fibroblast growth factor and vascular endothelial growth factor on subcutaneously allotransplanted ovarian tissue in ovariectomized mice. PLoS One 2015;10(7):e0134035. [44] Youm HW, Lee J, Kim EJ, Kong HS, Lee JR, Suh CS, Kim SH. Effects of angiopoietin-2 on transplanted mouse ovarian tissue. PLoS One 2016;11(11):e0166782. [45] Shiroma ME, Botelho NM, Damous LL, Baracat EC, Soares-Jr JM. Melatonin influence in ovary transplantation: systematic review. J Ovarian Res 2016;9(1):33.

[46] Cohen Y, Dafni H, Avni R, Fellus L, Bochner F, Rotkopf R, et al. Genetic and pharmacological modulation of Akt-1 for improving ovarian graft revascularization in a mouse model. Biol Reprod 2016;94(1):14. [47] Wang Y, Chang Q, Sun J, Dang L, Ma W, Hei C, et al. Effects of HMG on revascularization and follicular survival in heterotopic autotransplants of mouse ovarian tissue. Reprod Biomed Online 2012;24(6):646–53. [48] Abir R, Fisch B, Jessel S, Felz C, Ben-Haroush A, Orvieto R. Improving posttransplantation survival of human ovarian tissue by treating the host and graft. Fertil Steril 2011;95(4):1205–10. [49] Fabbri R, Sapone A, Paolini M, Vivarelli F, Franchi P, Lucarini M, et al. Effects of N-acetylcysteine on human ovarian tissue preservation undergoing cryopreservation procedure. Histol Histopathol 2015;30(6):725–35. [50] Sánchez-Serrano M, Novella-Maestre E, Roselló-Sastre E, Camarasa N, Teruel J, Pellicer A. Malignant cells are not found in ovarian cortex from breast cancer patients undergoing ovarian cortex cryopreservation. Hum Reprod 2009;24:2238–43. [51] Azem F, Hasson J, Ben-Yosef D, Kossoy N, Cohen T, Almog B, Amit A, Lessing JB, Lifschitz-Mercer B. Histologic evaluation of fresh human ovarian tissue before cryopreservation. Int J Gynecol Pathol 2010;29:19–23. [52] Dolmans MM, Iwahara Y, Donnez J, Soares M, Vaerman JL, Amorim CA, Poirel H. Evaluation of minimal disseminated disease in cryopreserved ovarian tissue from bone and soft tissue sarcoma patients. Hum Reprod 2016;31:2292–302. [53] Rosendahl M, Timmermans Wielenga V, Nedergaard L, Kristensen SG, Ernst E, Rasmussen PE, et al. Cryopreservation of ovarian tissue for fertility preservation: no evidence of malignant cell contamination in ovarian tissue from patients with breast cancer. Fertil Steril 2011;95:2158–61. [54] Abir R, Aviram A, Feinmesser M, Stein J, Yaniv I, Parnes D, et al. Ovarian minimal residual disease in chronic myeloid leukaemia. Reprod Biomed Online 2014;28(2):255–60. [55] Nichols-Burns SM, Lotz L, Schneider H, Adamek E, Daniel C, Stief A, et al. Preliminary observations on whole-ovary xenotransplantation as an experimental model for fertility preservation. Reprod Biomed Online 2014;29:621–6. [56] Kawahara T, Sugishita Y, Taylan E, Suzuki N, Moy F, Oktay K. Vitrification versus slow freezing of human ovarian tissue: a comparison of follicle survival and DNA damage. Fertil Steril 2017;108: e56–7. [57] Sugishita Y, Taylan E, Kawahara T, Suzuki N, Oktay K. Comparison of open and closed devices in human ovarian tissue vitrification. Fertil Steril 2017;108:e172–3.

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C H A P T E R

31 Oncofertility: Preservation of Ovarian Function After a Cancer Diagnosis Azza M. Akasha, Teresa K. Woodruff MINIMIZING THE IMPACT OF CANCER THERAPY ON THE OVARY

Abbreviations AAD AFC AMH ART CED COC DVT ESC FSH GnRHa IUD IVF PE S1P VTE

alkylating agent dose antral follicle count anti-M€ ullerian hormone assisted reproductive technology cyclophosphamide equivalent dose combined oral contraceptive deep vein thrombosis embryonic stem cell follicle-stimulating hormone gonadotropin-releasing hormone agonist intrauterine device in vitro fertilization pulmonary embolism sphingosine-1-phosphate venous thromboembolism

INTRODUCTION Advances in cancer diagnostics and treatment have led to a significant increase in survival rates, particularly in pediatric cancers, resulting in a paradigm shift in focus toward long-term survivorship and quality-of-life issues. While cardiac, pulmonary, renal, and liver functions are frequently assessed before, during, and after treatment, the same is not true for the reproductive system. As greater numbers of girls and women survive cancer, it is essential to anticipate the impact of cancer and its treatment on the ovary and offer options to protect the future reproductive and sexual function of this population. The term “oncofertility” was coined to bridge the gap between the disciplines of oncology and reproductive endocrinology and provide a common platform where clinicians, scientists, and scholars can provide fertility preservation options to patients diagnosed with cancer. This chapter reviews the risks to fertility posed by cancer therapies, how fertility is measured using markers of the ovarian reserve, current and experimental fertility preservation options, and contraceptive considerations for patients with cancer.

The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00031-5

Several options are available for fertility preservation after a cancer diagnosis, prior to initiation of cancer treatment. Oocyte and embryo banking are now standard of care, while ovarian tissue cryopreservation and immature oocyte cryopreservation are investigational techniques [1]. However, these interventions are invasive, costly, and may require a delay of cancer treatment for up to 2 weeks [2], which may not be an option for patients with aggressive disease (Table 1). Minimizing the effect of the planned cancer treatment on the ovaries may help preserve fertility and reproductive function. It is important for clinicians to clarify the risk of gonadal injury caused by the planned cancer treatment regimen and refer patients for fertility preservation consultation. Alkylating agents, in particular, have been shown to diminish the ovarian reserve [3,4]. Chemotherapy-related infertility risk can be estimated by calculating the cumulative dose of alkylating agents that will be received. Both the alkylating agent dose (AAD) and the cyclophosphamide equivalent dose (CED) can be used to quantify the risk of gonadal toxicity. To minimize risk of ovarian tissue damage during cancer treatment, several potentially fertoprotective agents have been developed. These agents aim to decrease the need for invasive, costly, and time-consuming fertility preservation therapies prior to the start of cancer treatment. These agents include the following: • Gonadotropin-releasing hormone agonist (GnRHa): Although frequently studied, the use of GnRHa remains controversial due to conflicting results on long-term ovarian function and pregnancy outcomes. GnRHa does not interfere with cytotoxic treatments and have been used in the clinical setting [5–7].

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502 TABLE 1

31. ONCOFERTILITY: PRESERVATION OF OVARIAN FUNCTION AFTER A CANCER DIAGNOSIS

Effects of cancer therapy in female fertility

Treatment administered

Cause or disease

Adverse effects of treatment

Mechanism

Surgery

Ovarian tumor (late stage)

Impairs fertility Impairs hormone production

Decreases number of follicles present

Cystectomy/unilateral oophorectomy

Ovarian tumor (early stage)

None

Remaining ovary compensates when no postoperative chemotherapy or radiation are given

Pelvic surgery

Nongynecological malignancies

Impaired folliculogenesis Impaired ovulation Impaired tubal transport

Cytokine production Formation of pelvic adhesions

Chemotherapeutic agents

Malignancies

Temporary amenorrhea Primary ovarian insufficiency

DNA damage to mature follicles with subsequent apoptosis Decreases reserve primordial pool

Radiotherapy dose dependent

Malignancies

Infertility due to ovarian, uterine, hypothalamus, or pituitary damage

Radiation doses greater than 2 Gy result in 50% loss of ovarian follicles Irreversible damage to uterus in abdominopelvic doses greater than 30 Gy Radiation to hypothalamus/pituitary greater than 30 Gy affects gonadotropin production

• Imatinib: A competitive tyrosine kinase inhibitor that blocks apoptosis by inhibiting c-Abl-mediated upregulation of tumor suppressor p63. Imatinib reduces primordial follicle loss when administered prior to cisplatin treatment [8]. Concerns remain as to whether spared oocytes can harbor DNA damage, which may result in miscarriages or birth defects [9,10]. • Sphingosine-1-phosphate (S1P): Regulates proliferative cellular processes including inhibiting apoptosis. Studies on mice have been inconclusive with regard to the efficacy of S1P as a fertoprotective agent; some studies have shown a protective effect in the presence of dacarbazine, while others have shown no effect on the presence of cyclophosphamide [11]. S1P is administered by tissue injection rather than systemic administration, which limits its clinical use. • Tamoxifen: Although rodent studies have shown an ovarian protective role for tamoxifen prior to cyclophosphamide and radiation, human studies have shown no protection [12–14]. Inconsistencies in the data may be due to differences in study design and use of different endpoints, underscoring the need for welldesigned human studies for further investigation [15]. • AS101: A tellurium-based immunomodulatory agent shown to protect gonads against chemotherapyinduced follicular damage without interfering with cancer treatments. AS101 has also demonstrated an antitumor effect in rodents and human studies [16,17].

ASSESSMENT OF THE OVARIAN RESERVE The ovarian reserve, defined as the population of follicles that is capable of producing a mature egg for

fertilization, can be used as a measure of a woman’s fecundity, oocyte quality, and quantity. Most cancer survivors experience infertility due to the cytotoxic effect of cancer treatment on the ovaries, hypothalamus, pituitary, or uterus. Radiation and other gonadotoxic agents reduce the number of ovarian follicles by accelerating the process of attrition [18,19]. The ovarian reserve varies between individuals based on the age at the time of gonadotoxic treatment, the type and dose of therapy, genetic factors, previous illnesses, and prior infertility. Quantifying the ovarian reserve prior to starting cancer treatment may help predict the likelihood of future fertility identify those in need of fertility preservation procedures. Markers of the ovarian reserve include the following: 1. Menstrual cycle: The average age of menarche in the western world is 12 years, with a mean cycle interval of 32 days (range 21–45 days) and flow length of 7 days [20]. Absence of menses by the age of 15 is defined as primary amenorrhea, while secondary amenorrhea is the absence of already established menses for >6 months. The average age of menopause in the United States is 51 years, and cessation of menses before the age of 40 is considered early menopause. Adult female survivors of childhood cancers have been noted to have a higher rate of early menopause compared to the general population. Although traditionally used as the primary measure of fertility and ovarian function in cancer survivors, menses can be affected by many factors including hypogonadotropic hypogonadism and structural issues. Women may also continue to have regular menses despite a diminished ovarian reserve, making this marker a poor predictor of ovarian reserve.

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FERTILITY PRESERVATION OPTIONS IN THE CANCER SETTING

2. Antral follicle counts (AFCs): AFC utilizes ultrasound to assess the number of small follicles (2–9 mm) in both ovaries during the early follicular phase of the cycle [19]. Tests revealing low AFC correlate with poor response to ovarian stimulation, yet, cannot accurately predict future infertility. AFC demonstrates inter-cycle and inter-observer reliability, and thus can be considered a promising screening tool for assessing the ovarian reserve. 3. Ovarian volume: While ovarian volume can be utilized to estimate follicle count, it exhibits poor inter-cycle reliability and should be used in combination with AFC [19]. In children, assessment of AFC and ovarian volume can be performed by a skilled radiologist using transabdominal ultrasound. 4. Follicle-stimulating hormone (FSH), inhibin B, and estradiol: Consistently high levels of FSH are predictive of diminished ovarian reserve and levels above 40 IU/L are diagnostic for premature ovarian insufficiency or menopause [19]. Basal estradiol levels do not differ between women with and without diminished ovarian reserve. In prepubertal children, estradiol levels are suppressed by the hypothalamus [19]. Inhibin B secreted by the preantral follicles may be a good indicator of ovarian reserve. 5. Anti-M€ ullerian hormone (AMH): AMH reflects changes in ovarian function earlier than other markers such as inhibin B or estradiol. AMH does not significantly fluctuate during the menstrual cycle, and it is highly predictive of menopause timing [21,22]. AMH levels are also detectable in females from birth to menopause [23], making it a suitable marker of the ovarian reserve in prepubertal girls. Additionally, AMH levels were found to be lower in women treated with mechlorethamine, vincristine, procarbazine, prednisolone, and abdominal and total body irradiation compared to those not treated with these drugs and healthy women. In adult women with cancer, AMH declines during treatment, followed by recovery in some patients. The rate of AMH decline is determined by pretreatment AMH levels. Additional research is needed to determine how AMH can be used to guide fertility preservation treatment and predict long-term ovarian function after cancer therapy.

FERTILITY PRESERVATION OPTIONS IN THE CANCER SETTING Family building has been identified as an important component of survivorship [24,25], and many fertility preservation options are currently available to female patients with cancer. These options can be divided into

503

those that can be applied before cancer therapy starts and those that are available for women who have already been exposed to gonadotoxic therapies.

Before Cancer Therapy Embryo and Oocyte Cryopreservation Embryo and oocyte cryopreservation are considered standard of care options in postpubertal patients at risk of ovarian failure [26]. They require controlled ovarian stimulation, traditionally beginning on the third day of the menstrual cycle, although newer protocols have been developed that are independent of the menstrual cycle [27,28]. Ovulation is triggered by a single dose of human chorionic gonadotropin, and transvaginal oocyte retrieval is performed 34–36 h later. The number of total and mature oocytes retrieved, the oocyte maturity rate, and the fertilization rate have been reported to be similar in both random and conventional ovarian stimulation cycles [28]. Oocyte cryopreservation may be preferred by patients who do not have partners at the time of retrieval, who do not wish to use donor sperm, or have ethical/religious objections to embryo freezing [29]. Ovarian Tissue Cryopreservation and Transplantation Although ovarian tissue cryopreservation and transplantation strategies are still in the early stages of development, this fertility preservation option is gaining popularity as it remains the only one that is available to prepubertal females and postpubertal females who cannot undergo hormonal ovarian stimulation [30]. The procedure involves surgical removal of the ovary and dissection of the cortical tissue for cryopreservation. When fertility is desired, the ovarian tissue is thawed and transplanted orthotopically or heterotopically back into the patient. Transplanted follicles can then be maturated by appropriate hormonal stimulation. Maturation of immature follicles retrieved from ovarian tissue remains an area of active research as it would eliminate the need for autotransplantation and avoid reintroducing cancerous cells that may be present in the ovarian tissue [31].

After Cancer Therapy Donor Oocytes and Embryos Patients in acute ovarian failure secondary to cancer treatment who maintain an intact and functional uterus may consider the use of donor oocytes or embryos. Oocytes from donors are fertilized using the intended parent partner or donor sperm and transferred into the intended mother’s uterus. The intended mother receives hormonal medication to modify her cycle in preparation for the embryo transfer. Should pregnancy be achieved, hormonal treatment continues into the third month of

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31. ONCOFERTILITY: PRESERVATION OF OVARIAN FUNCTION AFTER A CANCER DIAGNOSIS

gestation; women without ovaries continue to receive exogenous hormones throughout the remainder of the pregnancy [32]. Gestational Carrier or Adoption The use of gestational carriers may be considered by patients who do not have a uterus, have uterine damage, or have another medical condition that would limit or prevent pregnancy. This option requires medical and psychological evaluation for all parties involved. In addition, legal council is required [33]. For patients who are unable to conceive biological children, adoption is an option for family building. Although adoption agencies require medical documentation of a prospective parent’s health status, many are supportive of cancer survivors and only ask for a physician’s note outlining health status posttreatment [34].

endocrine function and initiation of puberty in a mouse model were achieved by utilizing a decellularized matrix of an ovary and isolated ovarian cells [40]. The technique utilizes a biologically active ovary-specific extracellular matrix on which isolated follicles are seeded. Another method under development uses a bioengineered artificial ovary consisting of mouse ovarian follicles seeded within a three-dimensional (3D)-printed scaffold. The scaffold is modeled based on the ovarian extracellular matrix composition and architecture to create a niche in which follicles can reside and grow [41]. In the future, induced pluripotent stem cells derived from the patient could provide a source of endocrine cells or even oocytes for fertility preservation; human embryonic stem cells (ESCs) have been differentiated into both populations [42,43]. Stem cell-based techniques would theoretically provide an unlimited source of patient-specific cells to achieve the complete restoration of the ovary after cancer therapy.

Experimental Fertility Preservation Methods Ovarian tissue cryopreservation and in vitro follicle maturation are currently areas of high research interest, as they may be the only option for some patients with cancer to preserve their fertility. In prepubertal patients, primordial follicles retrieved from ovarian cortical tissues, encapsulated in alginate beads, and cultured to support the growth of follicles to the antral stage [35]. Success of this approach in several animal models and nonhuman primates suggests that cryopreservation of ovarian tissue for fertility preservation is a valid option and can serve populations in which egg/embryo banking is not an option. Transplantation of ovarian cortical strips has been shown to restore endocrine function in patients who receive gonadotoxic treatments. Under a research protocol, 4–5 cortical strips (1 cm
0.5 mm
1.5 mm) were isolated and cryopreserved prior to gonadotoxic treatment, then thawed and fixed onto the remaining portion of the ovary, or inserted underneath the cortical capsule or inserted under the skin of the forearm to restore some endocrine function or stimulate mature eggs for in vitro fertilization (IVF) after cancer remission [36–38]. There have been 60 reported live births from ovarian cortical tissue transplants with varying degrees of success [39]. This may be attributed to the type and degree of treatment received, the amount of ovarian tissue remaining prior to transplant, and the number of follicles transplanted within the strip. Patients should be made aware of the risks associated with ovarian tissue transplantation, such as the reintroduction of cancerous cells, as the tissue was harvested prior to treatment. To minimize this risk, recent work has focused on ways to restore ovarian function without transplantation of whole ovarian tissue. In one study, restoration of

CONTRACEPTION AND MENSTRUAL SUPPRESSION FOR ONCOLOGY PATIENTS Open and early discussions about contraceptive needs with patients undergoing cancer treatment are imperative to avoid unintended pregnancies [44]. Teratogenic exposure or pregnancy terminations can have a devastating impact on patients and their partners. It is also important to consider the efficacy and safety profile of each method and to take into account the patient’s medical history and lifestyle including technical, social, and religious factors. Contraceptive options that limit or suppress menses may provide added benefits to cancer patients with low blood count, menorrhagia, and/or those who are at risk of bone marrow suppression. The available methods of contraception, their efficacy, safety profiles, ease of use, side effects, and oncology concerns are outlined in Table 2. For cancer survivors, at least 6 months in remission, and without a history of chest wall radiation, hormonally mediated cancers, anemia, osteoporosis, or venous thromboembolism (VTE), any of the abovementioned methods can be used. Suppression of menstruation can be used to prevent anemia and thrombocytopenia. Patients with thrombocytopenia as a result of their malignancy are at increased risk for hemorrhage, so suppression of menses can be a beneficial side effect of contraception. Some pharmaceutical agents used to regulate or suppress menstruation include the following: • GnRHa: these drugs can be given intravenously, subcutaneously, or intramuscularly, with a frequency of administration varying according to route used.

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CONTRACEPTION AND MENSTRUAL SUPPRESSION FOR ONCOLOGY PATIENTS

TABLE 2

Contraception options (MEC: US Medical Eligibility Criteria)

Category

Contraception

Failure rate within first year (best use, common use, %)

Behavioral method

Abstinence

0, 0–85

–

–

Withdrawal

4, 22

46

–

Male condom

2, 18

43

Female condom

5, 21

41

Diaphragm

6, 12

57

Latex or spermicide sensitivities can be seen. Diaphragm should not be left in place for a combined duration of longer than 24 h due to rare risk of toxic shock syndrome.

Pills

0.3, 9

67

Increased risk of VTE Nausea/vomiting

Advantageous for bone health

Transdermal patch

0.3, 9

67

Transient skin reactions More BRB than with pills

Advantageous for bone health Not recommended for patients >90 kg

Transvaginal ring

0.3, 9

67

Headaches Vaginal wetness

May not have positive effect on bone health

Progesterone only pill

0.3, 9

67

Break through bleeding Headache Nausea Breast tenderness Acne

–

Injectable progesterone

0.2, 6

56

Irregular vaginal bleeding Amenorrhea Weight gain Transient decrease in bone mineral density

Not recommended in treatments resulting in osteopenia/ osteoperosis

Progesterone implants

0.05, 0.05

84

May cause unpredictable vaginal bleeding

Irregular bleeding pattern may not be the best choice in patients with anemic concerns

LNG IUD

0.2, 0.2

80

Chance for amenorrhea

Used for treatment of endometrial hyperplasia and low-grade cancer May be considered for select cancer patients on tamoxifen

LARC nonhormonal

Copper IUD

0.8, 0.6

78

May increase menstrual blood flow

First line for breast cancer patients Not recommended in patients with anemic concerns

Emergency contraception

Combined OCPs LNG methods ulipristal copper IUD

–

–

Irregular vaginal bleeding Abdominal pain Headache Some breast tenderness

There is no circumstance in which he risks outweigh the benefit of EC

Barrier method

Estrogen progesterone methods

Short- or intermediateacting progesterone methods

LARC progesterone based

Continued use at 1 year (%)

Adverse risks

Cancer-specific issues These methods are ineffective for patients undergoing cancer treatments. It is important to discuss and counsel more effective methods Condoms should be advised in ALL sexually active patients to decrease risk of STI transmission Diaphragm may be a viable option in hormone-sensitive cancer patients who cannot use copper IUD

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Should be avoided in cancer patients due to increased risk of VTE. Pills, patches and rings are noted as category 4 in the MEC due to increased risk of DVT, current breast cancer, malignant liver tumors. Classified as category 1 for gestational trophoblastic disease and category 2 for cervical cancer awaiting treatment.

506

•

•

•

•

31. ONCOFERTILITY: PRESERVATION OF OVARIAN FUNCTION AFTER A CANCER DIAGNOSIS

These drugs offer an amenorrhea rate of 73%–79% [45]. The main adverse effects of GnRHa are vasomotor symptoms, decrease in bone mineral density, and risk of local contusions and hematomas at the site of injection. Patients can be offered add-back therapies including norethindrone acetate or low-dose estradiol with progestin to prevent loss of bone density [46]. Progestin-only pills: High-dose progestin-only pills can be used in a suppressive method, with amenorrhea achieved in a short period of time. Low-dose progestin only daily pills can also be used to achieve amenorrhea at a rate of 10% [47]. Depot medroxyprogesterone acetate: Rates of amenorrhea with this method are about 50%, though initial irregular vaginal bleeding is common [48]. This may not be tolerated by patients who have or are at risk of having low blood count due to their malignancy or chemotherapy regimen. Combined oral contraceptives (COCs): When taken in extended regimens without the free intervals, 70% of women become amenorrheic after 1 year of use [49]. Patients may experience breakthrough bleeds that may not be well tolerated by those with low blood counts. Levonorgestrel intrauterine device (IUD): 50% of patients note suppression of menses after 24 months of use. If a patient has an established suppression with the IUD in place at the time of her cancer diagnosis, it is recommended that the IUD be left in place [50].

CONCLUSION Fertility preservation for patients with cancer requires further research and optimization to address both endocrine and fertility needs in all female populations, prepubertal to menopausal. Some experimental paradigms utilizing tissue extraction to house the ovarian follicle reserve in females are promising. Culture of isolated follicles could provide a potential gamete and the option of biological offspring following assisted reproductive technology (ART) procedures. A collaborative effort between reproductive biologists, endocrinologists, regenerative medicine scientists, clinicians, therapists, and engineers is necessary to not only provide future options for cancer patients with regard to their reproductive needs but also to provide comprehensive care that considers every aspect of patients’ lives.

Glossary Fertility preservation Preservation of reproductive function, often used by REIs to refer to IVF interventions with oocyte or embryo cryopreservation. Gonadotoxicity Iatrogenic effects of chemo- or radiation therapy on ovarian function

Oncofertility Oncofertility includes biological and nonbiological options for male and female cancer patients to preserve or restore fertility. Ovarian reserve The number of primordial follicles available for future fertility

Acknowledgments The authors thank Stacey Tobin and Chelsea Castleberry for editorial assistance.

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[40] Laronda MM, Jakus AE, Whelan KA, Wertheim JA, Shah RN, Woodruff TK. Initiation of puberty in mice following decellularized ovary transplant. Biomaterials 2015;50:20–9. https://doi. org/10.1016/j.biomaterials.2015.01.051. PMID: 25736492; PMCID: PMC4350019. [41] Laronda MM, Rutz AL, Xiao S, Whelan KA, Duncan FE, Roth EW, Woodruff TK, Shah RN. A bioprosthetic ovary created using 3D printed microporous scaffolds restores ovarian function in sterilized mice. Nat Commun 2017;8:15261. https://doi.org/10.1038/ ncomms15261. PMID: 28509899; PMCID: PMC5440811. [42] Sasaki K, Yokobayashi S, Nakamura T, Okamoto I, Yabuta Y, Kurimoto K, Ohta H, Moritoki Y, Iwatani C, Tsuchiya H, Nakamura S, Sekiguchi K, Sakuma T, Yamamoto T, Mori T, Woltjen K, Nakagawa M, Yamamoto T, Takahashi K, Yamanaka S, Saitou M. Robust in vitro induction of human germ cell fate from pluripotent stem cells. Cell Stem Cell 2015;17 (2):178–94. https://doi.org/10.1016/j.stem.2015.06.014. PMID: 26189426. [43] Lan CW, Chen MJ, Jan PS, Chen HF, Ho HN. Differentiation of human embryonic stem cells into functional ovarian granulosa-like cells. J Clin Endocrinol Metab 2013;98(9):3713–23. https://doi.org/ 10.1210/jc.2012-4302. PMID: 23884780. [44] WHO. Medical eligibility criteria for contraception use August 16, 2017. Available from: http://apps.who.int/iris/bitstream/10665/ 181468/1/9789241549158_eng.pdf?ua¼1. [45] Quaas AM, Ginsburg ES. Prevention and treatment of uterine bleeding in hematologic malignancy. Eur J Obstet Gynecol Reprod

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Biol 2007;134(1):3–8. https://doi.org/10.1016/j.ejogrb.2007.03.012. PMID: 17467142. Divasta AD, Laufer MR, Gordon CM. Bone density in adolescents treated with a GnRH agonist and add-back therapy for endometriosis. J Pediatr Adolesc Gynecol 2007;20(5):293–7. https://doi.org/10.1016/j.jpag.2007.04.008. PMID: 17868896; PMCID: PMC3195423. Black A, Francoeur D, Rowe T, Collins J, Miller D, Brown T, David M, Dunn S, Fisher WA, Fleming N, Fortin CA, Guilbert E, Hanvey L, Lalonde A, Miller R, Morris M, O’Grady T, Pymar H, Smith T, Henneberg E. Canadian contraception consensus. J Obstet Gynaecol Can 2004;26(2):143–56 158–74, PMID: 15115624. Hubacher D, Lopez L, Steiner MJ, Dorflinger L. Menstrual pattern changes from levonorgestrel subdermal implants and DMPA: systematic review and evidence-based comparisons. Contraception 2009;80(2):113–8. https://doi.org/10.1016/j.contraception.2009.02.008. PMID: 19631785. Miller L, Hughes JP. Continuous combination oral contraceptive pills to eliminate withdrawal bleeding: a randomized trial. Obstet Gynecol 2003;101(4):653–61 PMID: 12681866. Anon. Committee opinion no. 606: Options for prevention and management of heavy menstrual bleeding in adolescent patients undergoing cancer treatment. Obstet Gynecol 2014;124 (2 Pt 1):397–402. https://doi.org/10.1097/01.AOG.0000452745.44206.c3. PMID: 25050771.

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32 Ovarian Cancers: Their Varied Origins and Pathologically Implicated Microenvironment Yu Sun*, Nelly Auersperg† †

*Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences, Shanghai, P.R. China Department of Obstetrics and Gynecology, University of British Columbia, Vancouver, BC, Canada

Abbreviations ADAM17 AGCT CAA CAF CCC CCL2 CM CSC EC ECM EOC EpCAM ER ERK2 EV FABP GI tract GREL GROa HGSC IFNγ IL JAK LGSC lncRNA MC MMP MSC O-ADSC OSE SLCT SFRP2 STAT STIC TGF-β TME TNF-α

a disintegrin and metallopeptidase domain 17 adult granulosa cell tumor cancer-associated adipocyte cancer-associated fibroblast clear cell carcinoma chemokine CdC motif ligand 2 conditioned media cancer stem cell endometrioid carcinoma extracellular matrix epithelial ovarian cancer epithelial cell adhesion molecule estrogen receptor extracellular regulated kinase 2 extracellular vesicle fatty acid-binding protein gastrointestinal tract gonadal ridge epithelioid like growth regulated oncogene-a high-grade serous carcinoma interferon gamma interleukin Janus kinase low-grade serous carcinoma long noncoding RNA mucinous carcinoma matrix metalloproteinase mesenchymal stem cell omental adipose-derived stem cell ovarian surface epithelium Sertol-Leydig cell tumor secreted frizzled-related protein 2 signal transducer and activator of transcription serous tubal intraepithelial carcinoma TGF-transforming growth factor-β tumor microenvironment tumor necrosis factor alpha

The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00032-7

TNFSF15 uPA VEGF ZEB

tumor necrosis factor superfamily-15 urokinase plasminogen activator vascular endothelial growth factor zinc finger E-box-binding homeobox

INTRODUCTION Cancers of the ovary are the fourth most common cause of cancer-related deaths among women worldwide and the prime cause of death from gynecological malignancies [1]. Ovarian cancer incidence increases with age, with a sharp increase in the rate from the mid-40s. In general, the ageadjusted incidence rate for ovarian cancer among all women is nearly 12 cases per 100,000. However, women under 65 have an incidence rate of 7.5 cases per 100,000, while those 65 and older are subject to an incidence rate of >42 cases per 100,000. Women carrying BRCA mutations have a significantly higher incidence of ovarian cancer and develop the disease even at an earlier age [2]. In spite of intensive research and general advances in cancer treatment, the 5-year survival among women diagnosed with ovarian cancer has remained near 40% over several decades. This limited advance in clinical management is largely the result of problems particular to the ovary: first, there is no reliable method to discover ovarian cancers in their early stage, when they are still curable; in many instances, the mass of the tumor at the time of discovery is so large that the exact site of origin cannot be determined. Second, though many ovarian cancers tend to respond well to initial chemotherapy, a high proportion of them recur in a chemotherapyresistant form. Third, malignant ovarian tumors vary

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strikingly in terms of histology and molecular characteristics as well as clinically, by parameters such as different age groups, progression, responses to therapy and prognosis. This variability reflects the fact that these malignancies represent distinct subtypes of cancer, which greatly complicates clinical management [3, 4]. Interestingly, while the specific cells of origin of cancers have been well characterized in most tissues, it is still uncertain where all the subtypes of ovarian cancers arise from. In recent years, there have been major changes in our knowledge and interpretation of the origin of the cancers that are found in the human ovary. It used to be accepted that most cancers detected in or on the ovary are, in fact, of ovarian origin. In particular, the epithelial ovarian cancers (EOCs), which represent the most common subtypes and cause the most deaths, were all thought to originate from the ovarian surface epithelium (OSE), that is, the modified pelvic peritoneum overlying the ovary [5, 6] and, more specifically, from cortical inclusion cysts within the ovarian stroma, that are lined by internalized OSE [7]. Anatomically, OSE is part of the pelvic peritoneum and as such is commonly defined as a mesothelium. However, it, in fact, displays a mixed epithelial/mesothelial phenotype: it concurrently expresses epithelial markers such as the basement membrane components laminin and collagen IV, simple sesmosomes, and keratins 7, 8, 18, and 19, as well as the mesenchymal components calretinin, vimentin, and collagens I and III. It is therefore not terminally differentiated but multipotent, which formed the basis for the assumption that it could give rise to epithelial cancers. Gene expression profiling has confirmed that OSE is multipotential, expresses stem cell characteristics, and appears capable of acting as cancer-initiating cells [6]. Perhaps the most striking change in this view has been the current concept that many of the ovarian cancer subtypes actually do not originate in the ovary but, rather, metastasize to the ovary from other sites [8]. A corollary of this idea is that the ovarian stroma, where these neoplasms develop, appears to be a particularly favorable site for the enhancement of invasion and growth by a variety of metastatic cells [9, 10]. This concept implies that many “ovarian” cancers are ovarian only by location, but not by origin, in contrast to malignant tumors at most other sites, where the location of the primary tumor indicates its origin. For the sake of simplicity, they are still referred to as ovarian cancers in the present review. In this chapter, the subtypes of ovarian cancer are briefly described including new concepts regarding their origin, and the role of the ovarian stromal microenvironment in the growth and response to treatment of the neoplastic cells is discussed [11, 12]. This information is preceded by a very brief summary of the embryonic development of the ovary and the Mullerian ducts, which clarifies some basic aspects of the ovary’s pathology in the adult.

DEVELOPMENT OF THE OVARY AND THE MULLERIAN DUCTS N.B. This is a brief summary of issues relevant to this review; a separate detailed chapter on the development of the ovary can be found elsewhere in this volume. The development of the vertebrate urogenital system that comprises the kidneys, gonads, and urinary and reproductive tracts begins soon after gastrulation, through the differentiation of the intermediate mesoderm [13]. At the earliest stages of gonadal development, the gonads form as ridges on the ventromedial of the mesonephros (the second embryonic kidney). They are composed mainly of cells from two sources: from cells derived from the overlying mesonephric surface epithelium which, in turn, originates as part of the primitive coelomic epithelium that lines the coelomic cavity [14] and from mesenchymal cells derived from the mesonephros [15]. While their sex is determined genetically, the gonads are initially bipotential and undergo either male or female differentiation in response to the expression of a sequel of specific genes. Before sexual differentiation, mammalian embryos have two pairs of genital ducts: the Wolffian ducts, which form the male urogenital duct system and regress in the female embryos, and the paramesonephric (Mullerian) ducts, which are the primordia of the epithelia of the oviducts, uterus, and upper vagina and regress in male embryos. The Mullerian ducts originate under the influence of LIM1, a LIM class homeodomain protein [16], as invaginations of the mesonephric surface epithelium that overlies the anterior aspect of the mesonephros, adjacent to the coelomic epithelium overlying the ovary, that is, the future OSE [15] (Fig. 1). The developing Mullerian ducts are in intimate contact with the mesonephros, which plays an important role as an inducer and regulator of Mullerian duct differentiation [17]. An important set of closely related homeodomaincontaining transcription factors in Mullerian tract development are PAX2/PAX8, which are expressed in the Mullerian and Wolffian ducts as well as in the developing kidneys. Among other functions, there is evidence that during development, PAX2/PAX8 is required for mesenchyme-to-epithelial transitions, such as in the formation of kidney tubules from mesonephric and metanephric msenchyme. The lack of either of these transcription factors results in major defects in the development of the reproductive tracts [13]. In adults, PAX proteins play a role in stem cell regulation, apoptosis resistance, and repression of terminal differentiation [18]. In normal reproductive tissues of adult women, PAX8 is expressed by the secretory cells of Fallopian tubes, endometrial glands, OSE-lined ovarian cortical inclusion cysts, and variable numbers of OSE cells on the ovarian surface.(Fig. 2)

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FIG. 1 Dissected fetal mouse ovary and mesonephros complexes from the bipotential gonad up until the day of birth. (A) 11.5 dpc, (B) 12.5 dpc, (C) 13.5 dpc, (D) 14.5 dpc, (E) 16.5 dpc, and (F) day of birth D0. White arrow points to gonads, black arrow to mesonephros, white arrow head to M€ ullerian duct, and black arrow head to Wolffian duct. Scale bar ¼ 0.5mM. Note the close spacial relationship between the mesonephros and the gonad and the continuity between the mesonephric and the ovarian surface epithelium. Reproduced with permission from Fig. 3 in: Sarraj MA, Drummond AE. Mammalian foetal ovarian development: consequences for health and disease. Reproduction 2012; 143(2):151–163.

(A)

(B)

(C)

FIG. 2 Immunohistochemical demonstration of the transcription factor PAX8 in (A) Fallopian tube fimbrial epithelium, (B) OSE on the ovarian surface (the black arrow points at the monolayer representing the OSE), and (C) OSE-lined ovarian cortical inclusion cysts. The nuclei of the fimbrial epithelium and the inclusion cyst epithelia are uniformly positive, while the OSE on the ovarian surface is negative, suggesting that the inclusion cysts express PAX8 in response to stromal influences. PAX8 is thought to play a role in mesothelial/epithelial transdifferentiation. OSE, ovarian surface epithelium.

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SUBTYPES AND ORIGINS OF OVARIAN CANCER This review discusses the EOCs and the sex cordstromal ovarian tumors. Germ cell tumors, which make up about 3%–5% of ovarian cancers, have been described in detail [19] and are not included in this chapter.

The Ovarian Epithelial Cancers The epithelial cancers, which represent 85%–95% of ovarian carcinomas, are categorized further by their morphology, differentiation, and molecular profiles into highgrade serous (oviductal) (70%–74%), low-grade serous (3%–5%), endometrioid (10%), clear cell (10%–15%), and mucinous (2%–6%) carcinomas (Fig. 3). The EOCs have been defined as two groups on the basis of specific combinations of mutations and gene inactivation, which are associated with major differences in clinical characteristics [20]. Type I tumors include low-grade serous, low-grade endometrioid, clear cell, and mucinous carcinomas (MCs). These tumors are slow growing, frequently discovered at low stages and share lineages with preinvasive and benign precursors. Type I tumors are genetically more stable than type II tumors. Type II tumors include the high-grade serous, high-grade endometrioid, and undifferentiated carcinomas. These carcinomas are usually detected late and do not have well-defined precursors. Most common among these are the serous high-grade carcinomas, which account for most deaths associated with ovarian cancer. Type I and II tumors display specific mutations: KRAS, BRAF, and ERBB2 mutations occur in most low-grade serous carcinomas (LGSCs), whereas TP53 mutations are rare in these tumors. In contrast, the highgrade serous carcinomas (HGSCs), the predominant type II tumors, are highly unstable genetically, and are characterized by frequent mutations in TP53 and the sBRCA1, and BRCA2 tumor suppressor genes, but rarely contain the mutations found in type I tumors. Low-grade endometrioid carcinomas (ECs) have mutations of CTNNB1, PTEN, and PIK3CA, while most MCs have KRAS

mutations and clear cell carcinomas (CCCs) exhibit PIK3CA-activating mutations. As mentioned above, the progenitor of all epithelial ovarian carcinomas was thought for many decades to be the OSE, a specialized part of the pelvic peritoneum which covers the ovary [5]. During embryonic development, the OSE originates as part of the multipotential coelomic epithelium, a mesothelium that covers the embryonic coelomic cavity and is the precursor of the mesonephric mesothelium and OSE, the pleura and peritoneum, as well as of many important embryonic structures such as the sex cords (ovigerous cords) and the Mullerian ducts. The Mullerian ducts develop from invaginations of the coelomic epithelium overlying the anterior part of the mesonephros, which is contiguous with the region of the coelomic epithelium that develops into the OSE (Fig. 1). This spatial relationship suggests that the site of origin of the Mullerian ducts in the mesonephric surface epithelium and the adjacent OSE are overlapping embryonic fields with overlapping potentials to differentiate. In the adult, the OSE overlies the ovary and lines surface invaginations as well as OSElined inclusion cysts within the stroma (Fig. 2). OSE is pluripotential as indicated by its stem cell characteristics [21, 22] and its propensity to undergo metaplasia toward stromal [23] or epithelial [24, 25] phenotypes, depending on environmental conditions and hormonal influences. The idea that EOCs originate in the OSE was originally introduced by Scully, who described lesions in the ovarian stroma that appeared to arise in OSE-lined inclusion cysts and resembled early serous carcinomas [26]. This concept was subsequently expanded to encompass also endometrioid and MCs, based on the fact that the mucosa of the fallopian tube, the endometrium, and the mucous epithelium of the cervical canal are all derived embryologically from the Mullerian duct, which originates in the same embryonic field as OSE [14, 15]. Much evidence indicates that human OSE as well as OSE from other species has the potential to undergo neoplastic transformation to neoplasms resembling ovarian carcinomas in a variety of aspects [27–29]. Two recent papers used mouse ovaries as their research model and identified ovarian

FIG. 3

The subtypes of ovarian malignant tumors and their relative frequencies.

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stem/progenitor cells in the OSE [30, 31]. In the study by Ng et al., these cells were identified using the expression of Lgr5, a stem cell marker of multiple epithelia. In adult mice, Lgr5 was expressed within proliferative regions of the OSE and at the junctions between the epithelia of mesovarian ligaments and the oviductal fimbriae, thus marking stem/progenitor cells of the ovary and tubal epithelia [31]. These results support the concept that zones of transition between different cell types exhibit increased susceptibility to malignant transformation [32]. However, in recent years, evidence has accumulated indicating that many subtypes of EOC arise at sites outside the ovary and can be considered as metastases to the ovary. Whether these are the sole sites of origin of EOC or not is still under investigation. Taken together, these data suggest that many of the “ovarian” neoplasms are derived from extraovarian sources and raise the interesting and important question why the ovarian stroma is a preferential site for metastases of so many distantly originating neoplasms. This question is discussed in the second part of this chapter.

Subtypes of Ovarian Epithelial Cancers High-Grade Serous Carcinomas HGSCs are the most common type of ovarian cancer, accounting for roughly 70%–74% of all epithelial ovarian carcinomas, and are responsible for most deaths due to ovarian cancer. They are predominantly a disease of postmenopausal women; they do, however, also occur in premenopausal women, in particular, in those carrying BRCA1 mutations [8]. Fewer than 5% of HGSCs are diagnosed at Stage I, when the disease is confined to the ovaries. It is usually diagnosed at late stages, when the tumor has spread beyond the ovary to the omentum and other intra-abdominal locations, with or without ascites. At these stages, the rate of recoveries is very low. Serous ovarian carcinomas, by definition, exhibit serous, that is, oviductal epithelial differentiation. By histopathology they are characterized by papillary or solid growth with slit-like spaces, with the tumor cells characterized by abnormal cell nuclei, and by abundant cell proliferation (Fig. 4). Over 90% of HGSCs have mutations at TP53.

FIG. 4 Representative examples of the major histological types of epithelial ovarian carcinomas. Reproduced with permission of Annual Review of Pathology, Volume 4 by Annual Reviews, http://www.annualreviews. org. From Fig. 1 in: Cho KR, Shih IeM. Ovarian cancer. Annu. Rev. Pathol. 2009;4:287–313.

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Other common molecular changes include BRCA1 loss in 30%–45% of cases and chromosomal instability/aneuploidy in 100% of cases [33]. There is now considerable evidence that most of the HGSCs originate in the secretory cells of the mucosa of the fallopian tube [8]. HGSCs were thought to be derived from OSE until 2001, when Piek reported the frequent presence in the oviductal fimbrial mucosa of small noninvasive lesions, which shared characteristics of serous carcinomas [34]. These lesions were particularly frequent in the fallopian tubes of women carrying BRCA1 mutations, a group with a high incidence of serous-type ovarian cancer. Piek proposed that cells, detached from these lesions, were deposited on the ovarian surface by the oviductal fimbriae at the time of ovulation, and were the precursors of the serous ovarian carcinomas. This discovery has since been investigated in great detail, and it has been confirmed that the majority of advanced high-grade serous ovarian cancers, currently estimated at about 60% [33], seem to be indeed derived from the oviductal epithelial lesions described by Piek. The oviductal lesions were subsequently named serous tubal intraepithelial carcinomas (STICs) [35]. It is important to note that STICs are rarely found in 100% of HGSC cases in any one study, suggesting that there is another source of these cancers. It has been proposed that this alternative source is endosalpingiosis that occurs when fragments of fimbrial epithelium detach and enter the ovarian stroma, perhaps via the ovulatory defect. Indeed, small cysts lined with benign tubal-type epithelium are often identified in the ovarian stroma, but it remains unclear whether and how this ectopic tubal epithelium is actually imported into the ovary. If the point of entry was really ovulatory defects, it would be expected that inclusion cysts or epithelial fragments lined by tubal epithelium would be found most frequently near corpora lutea or corpora albicans. But no such special relationship has been reported. Furthermore, there is to date no convincing evidence that the epithelium of these cysts undergoes (pre)neoplastic changes. Alternatively, there is evidence that neoplastic transformation may take place in OSE fragments that have translocated into the ovarian stroma through conformational changes of the ovarian surface and formed OSE-lined inclusion cysts. Such cysts undergo tubal metaplasia by losing mesenchymal characteristics such as calretinin and acquiring tubal epithelial characteristics such as PAX8, cilia, oviductspecific glycoprotein1, and epithelial cell adhesion molecule (Epcam) and may coexpress calretinin and PAX8 during this process [24, 36]. These characteristics imply that adult OSE remains in a multipotential progenitor state and is capable of acquiring Mullerian characteristics [24]. The current consensus is that about 60% of the HGSCs originate as STICs in the epithelium of the

oviductal fimbriae [33]. The origin of the remaining cases is still under dispute. Among ovarian neoplasms, the transcription factors PAX2/8 are found in serous, endometrioid, and CCCs [37, 38]. In a mouse model, PAX2 was found to act as a tumor suppressor or oncogene, depending on the model system [38]. PAX8 is an important determinant of mesenchyme-to-epithelial conversion during development, [13] and it has been used extensively in recent years as a marker of epithelial differentiation to determine whether HGSCs originate in the mesothelial OSE lining the ovarian surface and OSE-lined cortical inclusion cysts, or in the epithelial lining of the Fallopian tube fimbriae and inclusion cysts lined with tubal-type epithelium [8]. These investigations were based on the observation that the secretory cells of Fallopian tubes are consistently PAX8 positive, while OSE was considered to be PAX8 negative on the basis of a high proportion of ovaries with few or no PAX8-positive OSE cells lining the ovarian surface. This transcription factor was therefore used to argue that OSE was not a source of HGSCs. However, in a series of recent publications, PAX8 was again absent or only variably present in OSE on the ovarian surface, but was expressed by up to 100% of OSE cells in OSE-lined inclusion cysts [36, 37, 39, 40] (Fig. 2). Thus, PAX8 expression by OSE is induced or enhanced when the cells are translocated from the ovarian surface where they are isolated from the stroma by a collagenous tunica albuginea to the ovarian interior where they are exposed to stromal influences. Importantly, during development, the embryonic mesonephros is required to induce Mullerian duct differentiation, and mesonephric stromal cells migrate from the mesonephros into the fetal ovary [41] and presumably form part of the ovarian stroma. These processes suggest that the OSE-lined inclusion cysts express PAX8 in response to stromal inductive influences and that this is a step in the induction of OSE to undergo mesothelialto-epithelial metaplasia and, as a consequence, to express epithelial-type phenotypes in the case of neoplastic transformation. Low-Grade Serous Carcinoma LGSCs are uncommon; they comprise about 3% of all EOCs. They are relatively slow growing, and they tend to be more chemoresistant than other subtypes of ovarian cancer [42]. LGSCs are only rarely diagnosed at early stages and if they are diagnosed at late stages, their overall survival rate is poor. Histologically, LGSCs are characterized by papillary structures, low mitotic activity, and relatively uniform, small nuclei (Fig. 4). They frequently metastasize as small solid nests of tumor cells or as micropapillae. Despite the similarity in names, LGSCs rarely progress to HGSCs. The two types of ovarian carcinomas have different patterns of mutations: characteristic

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mutations in LGSCs include the mutually exclusive activating mutations of KRAS or BRAF and activating mutations of ERBB2. They tend to be genomically stable and diploid or near diploid and to express estrogen and progesterone receptors. There is clinical, genetic, and genomic evidence indicating that LGSCs evolve in a stepwise fashion from ovarian epithelial inclusions to benign serous cystadenomas, to borderline serous tumors, to serous tumors of low malignant potential (LMP serous tumors), and eventually to LGSCs [43]. Endometrioid Carcinoma ECs represent approximately 10% of all ovarian epithelial cancers. Patients with Stage I ECs tend to have excellent outcomes, but the overall 5-year survival of patients presenting with EC at higher stages is poor. By histopathology, ECs exhibit glandular formations resembling ECs of the uterus (Fig. 4). A large body of evidence indicates that ovarian ECs are derived from endometriosis, that is, implants of benign endometrial tissue (glands and stroma) outside the uterine cavity. Endometriosis is a common condition affecting women of reproductive age. It is thought to be most likely caused by retrograde menstruation where fragments of endometrial stroma and epithelium pass through the fallopian tubes into the pelvic cavity and implant on surrounding tissues, most commonly the ovaries. Here they form blood-filled cysts, known as endometriomas. That these structures are precursors of ECs is indicated by histologic [44] and molecular [45] evidence. Epidemiologic studies have consistently found two- to threefold increases in the risk of ovarian ECs in patients with endometriosis. Low-grade ECs often arise from endometrioid borderline tumors, which in turn may arise from endometriosis. This stepwise histopathological progression of ECs is frequently accompanied by the accumulation of mutations predicted to deregulate canonical Wnt/β-catenin/Tcf (Wnt) signaling and PI3K/PTEN signaling. Genes that are characteristically mutated in EC include CTNNB1, PIK3CA, KRAS, ARID1A, PTEN, and PPP2R1A. CTNNB1 mutations are very common in ECs but rare in all of the other major ovarian carcinoma subtypes. In contrast, genes such as PIK3CA and ARID1A are frequently mutated in both ECs and CCCs. Loss of PTEN function by LOH or mutation is an early event in the development of endometriosis-related cancers of the ovary. These mutations were found in contiguous atypical endometriosis but not in distant endometriosis, further supporting that local transformation of endometriosis tissue occurs. Taken together, the data strongly support a model for early mutation of ARID1A during malignant transformation of endometriosis. Thus, EC is one of the major types of ovarian carcinoma with a well-defined association

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with a benign extraovarian precursor. Interestingly, endometriotic lesions are found at several sites throughout the pelvis, but only those located on the ovary tend to progress to carcinomas. This suggests that the ovarian microenvironment is required for malignant progression to take place. Clear Cell Carcinoma CCCs make up 5%–25% of all ovarian cancers, making them the most common type after the HGSCs [46]. They are so named because the cancer cells typically have abundant clear cytoplasm due to the presence of intracytoplasmic glycogen, which is lost during processing (Fig. 4). Nearly half of CCCs are diagnosed at Stage I; yet, the overall prognosis is relatively unfavorable because at late stages the prognosis is worse than that of all other types of ovarian cancer at similar stages, as a result of resistance to standard chemotherapy and recurrent disease. Besides ECs, CCC is the other major ovarian carcinoma that is causally associated with endometriosis. Growth patterns of CCCs vary (e.g., solid, papillary, and tubulocystic), with many CCCs arising in association with endometriotic cysts or benign tumors known as clear cell adenofibromas. Like ECs, a high proportion of CCCs contain mutations in ARID1A and PIK3CA. CCCs also have mutations in PPP2R1A, PTEN, KRAS, and TP53, but at lower frequencies than ECs. In addition to the histologic evidence, the resemblance in genetic changes to endometrial cancers indicates that CCCs likely originate in endometriosisrelated precursors. Molecular biology and genetics studies have shown that CCCs are usually negative for BRCA1 and BRCA2 mutations, but positive for ARID1A and PIK3CA mutations. The high frequency of PIK3CA mutations in CCCs (40%) leads to higher activity of the PI3K-AKT-mTOR pathway. Recent microarray experiments have revealed that CCCs have a genomic expression pattern that differs from serous EOCs, suggesting that the biologic phenotype of the clear-cell subtype is unique [46, 47]. Mucinous Carcinoma MCs are the least common of the major types of ovarian carcinoma, comprising about 3% of all EOCs. MCs remain problematic because there are still no clear indicators to distinguish borderline from invasive mucinous ovarian tumors and intrinsic cancers from metastatic ones. While the prognosis for MCs of all stages combined is relatively good, and patients with Stage I disease at diagnosis have an excellent prognosis, the prognosis for patients with late stage disease is very poor with their overall survival significantly less than that of women with advanced serous carcinoma. The basis for this difference has not yet been defined.

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This group of tumors is characterized morphologically by a glandular architecture and stratified columnar cells with basally located nuclei and pale-staining mucin in the cytoplasm (Fig. 4). The cytoplasm tends to become mucin-depleted in high-grade MCs. KRAS mutations are found in 75% of MCs and TP53 mutations in roughly half of invasive MCs. These mutations frequently occur in the same tumors. ERBB2 amplification is found in 19% of MCs. ERBB2 amplification and overexpression and KRAS mutations are largely mutually exclusive, with either being present in about 65% of cases. Within MCs, either HER2 amplification/overexpression or KRAS mutation is associated with a decreased likelihood of disease recurrence or death when compared to the cases with neither feature. Until the 1990s, it was generally accepted that MCs were derived from the OSE and had differentiated along the lines of the mucus-secreting cervical epithelium, which is derived from the Mullerian duct. Therefore, the assumed frequency of primary MCs was significantly higher than it is now accepted, since several studies have now shown that many ovarian MCs are not primary to the ovary but rather metastases from MCs that arose in other organs such as the gastrointestinal (GI) tract (colon, appendix, pancreas, stomach, and biliary tracts), endometrium, and endocevix. According to Zaino et al. [48], 57%–63% of ovarian MSs are metastases, thus reducing the likely frequency of primary mucinous adenocarcinomas to 1.5%. These conclusions are based on evidence such as the chemical composition of the mucins in ovarian MCs, which frequently resembles mucins of the gastrointestinal (GI) tract rather than those found in the cervical canal. However, specific criteria to distinguish primary from metastatic MCs are still not available.

Sex Cord-Stromal Tumors Adult Granulosa Cell Tumors Granulosa cells surround the germ cells, nurture them, protect them at the time of ovulation, and produce hormones. Developmentally, they derive from the sex cords (ovigerous cords), which, until recently, were thought to be outgrowths of the OSE. This theory has recently been modified by the description of a new cell type: gonadal ridge epithelioid like (GREL), which is derived from the mesonephric surface epithelium and gives rise to two cells types: it proliferates to contribute to the formation of the gonadal ridge/ovarian primordium, which includes GREL-derived ovigerous cords surrounding the germ cells; the other GREL-derived cell type remains on the ovarian surface and gives rise to the OSE [14]. The germ cells, which originate in the yolk sac, migrate into the genital ridges where they differentiate into oocytes. The sex cords/ovigerious cords, each of which surrounds many germ cells, are eventually separated by ovarian

stroma into individual primordial follicles, composed of single germ cells covered by OSE/GREL-derived follicular cells. The stroma has two sources: OSE cells that underwent epithelial-mesenchymal transition, and mesenchymal cells that migrate into the genital ridges from the mesonephros [41]. At puberty, the follicular cells undergo growth, differentiation and multilayering and become functional steroidogenic granulosa cells. The adult granulosa cell tumors (AGCTs) tend to remain confined to the ovary and show morphologic, immunologic, and endocrinologic resemblances to normal granulosa cells [8]. They are slow-growing tumors and have a tendency for late recurrence. Almost all AGCTs have identical missense mutations in the FOXL2 gene [49]. FOXL2 is one of the earliest markers in ovarian development and regulates granulosa cell differentiation and function in the adult. The precise mechanism by which this mutation causes cancer is unknown, but it may involve the transforming growth factor-β (TGF-β) pathway, which regulates granulosa cell biology. Sertoli-Leydig Cell Tumors Sertoli-leydig cell tumors (SLCTs) are rare tumors that originate in granulosa cells of young to middle-aged women. They are frequently characterized by androgenic symptoms that are probably the result of mutations in developing granulosa cells that lead to a switch in differentiation pathways from female granulosa cells to male Sertoli cells [8]. A major role in the development of these tumors is that one allele of DICER1 is truncated in the germ line [50]. It has been suggested that the DICER1 mutations disrupt the balance between sex-determining signals in the development and/or maintenance of gonadal development to cause a female-to-male phenotypic switch. It is possible that this combination of mutations does not completely inactivate DICER1 but instead skews microRNA (miRNA) processing. A second specific hypomorphic missense mutation appears to be required for actual tumor development. Why these changes are oncogenic, and how they lead to tumors that reflect a very unusual cell fate decision is not known.

CONCLUSIONS AND QUESTIONS ARISING Many advances in our understanding of ovarian cancers have taken place in recent years. An important discovery was that many more tumor types than were previously assumed to arise in the ovary are in fact, at least in part, metastases from a variety of other organs. These include the HGSCs, endometrioid and CCCs, MCs and, perhaps, LGSCs. The discovery that a high proportion of the HGSCs, the most lethal of ovarian cancers, appear to originate in the fimbriae of the fallopian tube

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and metastasize to the ovary has been a major advance in our understanding of the etiology of this disease. In addition, this information has initiated a widespread effort to investigate the feasibility of prophylactic removal of the fallopian tubes as a means to decrease the number of cases of HGSCs. Particular emphasis has been placed on women at high risk of developing HGSCs because of BRCA mutations. Numerous clinical trials of this approach are ongoing, but it will take some time until definitive conclusions regarding the effectiveness of prophylactic salpingectomies as a means to prevent HGSCs will be established. The striking predominance of metastatic tumors in the ovary indicates that the ovarian stroma provides a particularly favorable environment for the growth of malignant cells from many different sites; examples are that STICS, the precursors of HGSCs, grow exuberantly within the ovary but only to a very limited extent at their natural site within the tubal epithelium; that endometrial fragments, thought to be the source of the ovarian endometrioidui cancers, settle in various sites within the pelvis, but preferentially progress to cancers in the ovary. But related questions and problems remain unsolved: these include the nature of the metastasis-friendly ovarian stroma and the reasons for the high incidence of secondary drug resistance, which is one of the major problems in clinical ovarian cancer management. The second half of this review deals with these two problems.

THE OVARIAN CANCER MICROENVIRONMENT: A PRELIMINARY OVERVIEW One of the main facts that cause difficulty in understanding the biology and evolution of ovarian cancer is that most cancer cells are phenotypically not similar to normal ovarian cells. For HGSC, the most common ovarian malignancy usually diagnosed at an advanced stage with high mortality, no credible precursor lesion was histologically identified until 15 years ago, with the majority of mucinous ovarian cancers being metastases from other organs including the pancreas [51]. Indeed, a wide variety of cancers that are not derived from normal ovarian cell types including metastatic cancers from the breast, lung, and the gastrointestinal tract (GI tract) exist as major ovarian masses. In contrast to the ovary, which is generally understood as a fertile environment supportive of precancerous and cancerous lesions, the fallopian tube acts as a presumably inhospitable microenvironment for growth of both primary and metastatic tumors [52, 53]. It is reasonable to speculate that the fallopian tube has evolved a tumorsuppressive microenvironment to minimize the chance of ectopic pregnancy, a condition essentially lethal until the era of surgery [8]. Of note, the size of most ovarian

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malignancies, including primary tumors and nongynecological metastases, is significantly larger than the normal ovary and among the largest malignancies in the body. Although tumor stroma is usually generated de novo, the pathophysiological role of the stroma in ovarian neoplastic growth remains unclear. Earlier studies indicated that ovarian tumor stroma displays both endocrinological and morphological features of normal adult ovary, while ovarian metastases from GI tract malignancies have stroma similar to neither that of the primary GI tract tumors nor extra-ovarian metastases by containing luteinized, steroidogenic ovarian stromal cells [54]. Thus, the stroma of ovarian tumors histologically resembles that of normal adult ovary, and can likely provide a microenvironment that promotes the development of primary and metastatic tumors. Beyond this, comparative profiling of the transcriptomics and proteomics of ovarian tumor stroma by contrasting with those of alternative tumor types such as primary breast cancer versus ovarian metastases may disclose stimulatory factors correlated with the preferential growth of ovarian cancer as potential therapeutic targets. Although it is well accepted that EOC cells are responsive to steroid hormone stimulation, new studies have presented the clues that the ovarian stroma may also have an active role in this process. For instance, ovarian stroma immediately adjacent to the tumor foci can express markers associated with sex-steroid differentiation and steroidogenesis (calretinin, inhibin, and steroidogenic factor 1), alongside steroid enzymes (CYP17, CYP19, HSD17β1, and AKR1C3), while the epithelium expresses corresponding hormone receptors [55]. Thus, the epithelium-surrounding stroma in the ovary is activated to elaborate biologically relevant hormones, which may enhance incontrollable neoplastic growth, although the precise mechanisms underlying these processes await further investigation. Specifically, isoform-specific alterations of Akt, the serine-threonine kinase whose three isoforms are encoded by distinct genes and frequently overexpressed in numerous cancers, was recently found to have divergent effects in ovarian cancer cells and the nearby microenvironment [56]. Ablation of Akt1 in the tumor microenvironment (TME) generated an inhibitory effect on tumor size, without significant change in animal survival, while elimination of Akt2 or Akt3 resulted in increased tumor size, metastasis, and decreased survival time [56]. Although it is increasingly evident that stromal components have significant clinical implications in ovarian cancer development, recent findings uncovered an even stronger impact orchestrated by diverse cell types that may predict overall and progression-free survival of HGSC [57]. Furthermore, quantitative histology-based assessments can further enable appropriate selection of patients who are in urgent need of specific therapeutic strategies including combinatorial treatments that target the heterogeneous TME [57]

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MULTIFACETED FUNCTIONS OF THE OVARIAN CANCER MICROENVIRONMENT IN DISEASE PROGRESSION A typical TME comprises diverse noncancerous cell lineages, including stromal fibroblasts, infiltrating leukocytes, adipocytes, neuroendocrine cells, endothelial cells (ECs), and pericytes [58]. According to the specific stage of disease progression and the particular organ type, TME cells can play tumor-promoting or tumorsuppressing roles, partially depending on the adjacent cancer cells that have coevolved. Importantly, some of the functional mechanisms through which the TME influences pathological progression are also “coopted” to drive ectopic metastasis and therapeutic resistance in clinical settings [59]. One of the main properties that distinguish ovarian cancer from other solid tumors is the specific TME within the ovary. As ovarian cancer is a peritoneal malignancy, cancer cell dissemination is partially dependent on the peritoneal fluid as a carrier [60]. In such a case, transcelomic dissemination is a major route of cancer cell adhesion to the omentum and serous membranes that line the peritoneal organs, generating metastatic lesions in the peritoneal cavity instead of invading through the peritoneal lamina propria [61]. The peritoneal environment is frequently formed by the effusion accumulating in the peritoneal cavity, which presents as large volumes of ascites. Typically, the ascites comprises detached cancer cells, numerous soluble factors, extracellular vesicles (EVs), various types of immune cells including T cells and tumor-associated macrophages, along with many other cell subpopulations, together favoring cancer cell proliferation, chemoresistance, and metastasis [62]. Distinct from most other human malignancies, metastases at distant sites are often confined to late stages of ovarian cancer, and the most serious problem for HGSC patients is recurrent and aggressive growth of metastatic lesions within the peritoneal cavity [62]. The second feature of ovarian cancer is the special relevance of the omentum, a physical structure composed of connective and fatty tissue that covers the ventral surface of the intestines. Specifically, the omentum is often the preferred site for ovarian cancer metastases and plays a key role in disease progression [61].

Cancer-Associated Fibroblasts Pathological development of ovarian cancer, from cell transformation to local tissue invasion and distant metastatic dissemination, relies on mutual communication between EOC cells and their adjacent stromal microenvironment. An appropriate understanding of the bidirectional interaction of early EOC cells with activated

stromal cells helps identify novel diagnostic stromal markers and molecular targets for clinical therapy. The stroma comprises up to 50% of the advanced ovarian tumor mass, wherein cancer-associated fibroblasts (CAFs) represent a major cell subpopulation in the local TME [63]. By producing secretory factors such as hepatocyte growth factor, CAFs remarkably decrease the sensitivity of cancer cells to various anticancer agents [64, 65]. Moreover, CAFs alter the tumor physical properties via excessive deposition and aberrant remodeling of the extracellular matrix (ECM), thus enhancing formation of an interstitial barrier that blocks efficient drug delivery [66]. Specifically, targeting CAFs increases bioavailability of chemotherapeutic agents including doxorubicin and gemcitabine, while enabling immunological surveillance and tumor destruction though a process that engages interferon-gamma (IFNγ) and tumor necrosis factor alpha (TNF-α) [67, 68]. The range of biological mechanisms employed by CAFs to mediate therapeutic resistance was expanded by a recent study, which identified a novel role of CAFs in minimizing cisplatin levels within ovarian cancer cells upon chemotherapy [69]. Specifically, CAF-derived glutathione and cysteine contribute to treatment resistance, which can be abolished by CD8(+) T cells by altering the metabolism of these molecules in CAFs [69]. CD8(+) T-cell-released IFNγ regulates glutathione and cysteine levels via upregulation of γ-glutamyl-transferases and transcriptional inhibition of system xc() cystine and glutamate antiporter through the Janus kinase/signal transducer and activator of transcription 1 (STAT1) axis. Importantly, the presence of stromal CAFs and CD8(+) T cells is negatively and positively correlated with the survival of ovarian cancer patients, respectively, thus capitalizing the interaction between chemotherapy and immunotherapy holds significant potential to improve treatment outcomes of ovarian cancer patients [69]. As immunotherapy is emerging as a mainstay of anticancer strategies in several malignancy types including ovarian cancer, novel avenues for therapeutically targeting the protumorigenic CAFs through modulation of CD8+ T cells are highly inspiring to both scientific and clinical communities. Although some technical issues still remain regarding how to effectively harness the TME in therapeutic settings, future studies that explore the interplay between the CAF populations and adaptive immune cells hold significant potential to provide updated approaches to integrate T cell therapy and/or fibroblast depletion, thereby enhancing the overall efficacy of DNA-damaging chemotherapies.

Mesenchymal Stem Cells MSCs represent an active stromal cell subpopulation that is recruited to the TME, with prominent

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multipotency that allows differentiation into various cell types. In certain cases, the recruitment may be partly induced by LL-37 (leucine, leucine-37), a proinflammatory peptide of human cationic antimicrobial protein 18, and other migration-promoting signals [70]. Human bone marrow-derived MSCs can differentiate into CAFs, which produce soluble protumorigenic factors such as interleukin 6 (IL-6) to enhance tumor growth in an EOC xenograft model [71]. Combining cancer cells and cancer-associated MSCs in vivo and in vitro causes activation of the bone morphogenetic protein signaling network, which plays important roles in cancer progression [72]. Coinjection of ovarian MSCs, which secret a high level (> 2500 pg/mL) of IL-6, with SKOV3 cells enhanced tumorigenesis, sphere and colony formation in nonobese diabetic-severe combined immunodeficiency mice, while administration of an IL-6 receptor blocking antibody minimized these malignant behaviors [73]. Interestingly, cancer exosome-treated MSCs have elevated α-smooth muscle actin expression, a change that indicates an activated fibroblast phenotype, alongside increased synthesis of tumor-promoting cytokines including stromal cell-derived factor-1 and TGF-β [74]. Thus, ovarian cancer-derived exosomes can contribute to the generation of CAFs differentiated from MSCs in tumor stroma. Development of enhanced chemoresistance to standard clinical therapies is not uncommon in cancer patients, frequently allowing cancer cells to acquiring a “cancer stem cell (CSC)-like” phenotype. This phenotypic change is usually accompanied by an EMT, the phenotypic switch mostly implicated in cancer metastasis. For instance, the metastatic cell line OVCA433 exhibits upregulated expression of EMT and stem cells markers including CD44, α2 integrin subunit, CD117, CD133, (EpCAM), Nanog and Oct-4, and enhanced activation of extracellular regulated kinase 2 (ERK2) signaling upon treatment with cisplatin [75]. To the contrary, ERK2 signaling blockage by a MEK inhibitor U012 diminished expression of EMT and CSC markers, implying the potential of targeting this pathway to reduce residual tumor burden, a common cause of ovarian cancer recurrence. Both proliferation and invasion of human EOC cells are remarkably enhanced upon coculture with omental adipose-derived MSCs (O-ADSCs) in vitro, and there is a global increase in protein expression in the EOC cells treated with conditioned media (CM) from O-ADSC [76]. Specifically, nine proteins were identified with differential expression after treatment with the CM, which are linked to carcinogenesis, apoptosis, and migration of cancer cells, suggest that O-ADSCs alter the proteomic profile of EOC cells via paracrine mechanism in favor of EOC progression [76].

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Ovary-Associated Adipocytes For years, adipocytes were considered as energy producing and storing residents of fat tissue in the TME. However, recent studies indicate that adipocytes may have other pathophysiological functions, and their interactions with cancer cells have been reported in breast, ovarian, colon, and gastric malignancies [77]. So far, tumor-promoting effects of adipocytes are linked to their secretion of adipokines, hormones, and growth factors including matrix metalloproteinase 11 (MMP11), IL-6, IL-1β into the surrounding TME, which collectively enhance the migration and invasion capacity of cancer cells [78]. In addition, upon coculture with breast cancer cells, adipocytes exhibit an activated phenotype characterized by enhanced production of proteases and cytokines including IL-6 and IL-1β, as well as delipidation and a loss of adipocyte-associated markers. Further, peritumoral adipocytes exhibit an altered phenotype with specific biological features that are sufficient to allow these cells to be named cancer-associated adipocytes (CAAs) [78]. Normal adipocytes stimulate the migration and invasion of cancer cells that are estrogen receptor (ER)-negative, a process mediated via a cytoskeletal element cofilin-1 and increased IL-6 secretion by adipocytes [79]. Beyond the adipose tissue, preadipocytes also reside in the bone marrow and stromal compartments of other organs such as skin [80]. Interactions between mesenchymal stroma and the underlying adipose tissue allow generation of MSCs and cytokines, each frequently triggering stromal cell senescence and increasing therapeutic resistance [81–83]. A new study exploring the mechanism of resistance to mitochondria-initiated apoptosis in EOC cells uncovered that adipocyte-induced upregulation of Bcl-xl in EOC cells was correlated with acquired chemoresistance, illustrating a novel pathway that allows the TME to modulate apoptosis-associated protein expression and confers chemoresistance on malignant cells [84]. A study using fluorescence technique to show the preferential migration of ovarian cancer cells to the mouse omentum revealed that cancer cell migratory behavior can be mediated by adipokines secreted by omentumassociated adipocytes, including chemokine CdC motif ligand 2 (CCL2), IL-6, IL-8, tissue inhibitor of metalloproteinase 1, and adiponectin [77]. Of note, coculture of adipocytes and ovarian cancer cells induced adipocytespecific lipolysis, allowing the transfer of free fatty acids to cancer cells, which in turn accelerated tumor cell proliferation through energy generation via β-oxidation. In addition, adipocyte-derived hormones such as leptin are associated with increased proliferation of ER-positive ovarian cancer cells, while ERα can be transcriptionally activated through the STAT3 signaling pathway, suggesting that both ER status and growth

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promoting properties of adipocytes need to be considered in obese ovarian cancer patients [85]. As adipokines can control multiple key processes including appetite, body temperature, blood clotting, energy expenditure, glucose homeostasis, inflammation, insulin sensitivity, reproduction, aging, and cancer [86], increasing studies begin to explore the functional roles of these circulating adipocyte-secreted factors in human cancer. To date, diverse adipokines such as adiponectin, leptin, resistin, plasminogen activator inhibitor-1 (PAI-1), vascular endothelial growth factor (VEGF), TNF-α, IL-6, autotaxin, and fatty acid-binding proteins (FABPs) are being extensively investigated [87]. Among them, adipose-derived FABP4 and FABP5 are identified as critical proteins particularly in lipid-related metabolic processes upon their overexpression in human malignancies including breast, prostate, colorectal, and ovarian cancers. For instance, CAAs interact metabolically with cancer cells through fatty acid transport via FABP4 in ovarian cancer, while FABP4 is involved in lipid transfer between adipocytes and cancer cells, a process that engages the fatty acid oxidation pathway to aid in cancer progression [88]. Interestingly, FABP4 knockdown in ECs causes increased fatty acid oxidation and reactive oxygen species generation, but decreased angiogenesis, growth, and metastasis in ovarian tumor xenografts [89]. Usually, high FABP4 expression levels in human primary tumors are correlated with elevated incidence of residual disease relapse after primary debulking surgery of HGSC, raising the possibility of exploring FABP4 as a candidate biomarker of residual disease in this condition [90].

Angiogenesis and Neovasculature In the TME, cancer cells rely on a constant supply of nutrients and oxygen, which is supported by the formation of new blood vessels. Several pathways are involved in the regulation of the growth and maintenance of neovasculature, a process that can be mediated by proangiogenic factors secreted by both cancer and stromal cells [91]. MMP1-mediated activation of the G protein coupled receptor and protease-activated receptor-1 stimulates ovarian cancer cells to release the chemokines CCL2, IL-8, growth-regulated oncogene-a (GROa) that induce EC proliferation, tube formation, and angiogenesis, to enable cancer cells to metastasize [92]. A new study examined vascular epithelial growth factor (VEGF) expression in benign, borderline, and malignant neoplasms to correlate it with histological grade and stage of ovarian cancer patients. Despite notable VEGF expression in some benign and borderline neoplasms, high VEGF expression was mostly observed in epithelial carcinomas, suggesting EOC as a candidate for VEGF-targeting therapy [93]. Although the globally

commercialized VEGF-specific antibody, Avastin, displayed effectiveness in suppressing angiogenesis in these animals, IL-8 and GROa-dependent endothelial tube formation in vitro remained unchanged [92]. Another study identified tumor necrosis factor superfamily-15 (TNFSF15), an endogenous suppressor of neovascularization that plays a critical role in the physiologically normal ovary but is lost in ovarian cancer [94]. TNFSF15 silencing before and after inoculation of ovarian cancer ID8 cells to mice markedly increases angiogenesis and tumor growth, suggesting that downregulation of TNFSF15 by cancer cells and tumor-infiltrating macrophages and lymphocytes is a prerequisite for neovascularization in ovarian cancer [94]. Pericytes, ECs, and smooth muscle cells are stromal components that play an essential role in angiogenesis. Although proangiogenic factors are produced by stromal and epithelial cells during early carcinogenesis or as a consequence of inflammation, enhanced angiogenesis is usually a late event in cancer progression [95]. Synthesis of the secreted frizzled-related protein 2 (SFRP2), a canonical modulator of Wnt signaling, is enhanced in damaged tumor stroma by chemotherapeutics, which promotes angiogenesis via a calcineurin/NFAT pathway in a noncanonical manner [96, 97]. Of note, SFRP2 acts as an agonist of WNT16B, another extracellular factor released by the therapy-damaged TME that significantly promotes drug resistance of residual cancer cells in human prostate, breast, and ovarian malignancies in the posttreatment stage [11, 97] (Fig. 5). Continuous accumulation of ascites, chronic inflammation, and elevated VEGF concentrations are among the typical hallmarks of ovarian cancer progression. Overexpressed c-Myc remarkably enhances VEGF concentrations in ascites of ovarian cancer in mice, while transduced KRAS significantly enhances inflammatory cytokine concentrations and increases the number of pericytes in animal ascites, suggesting that oncogenes can favor disease progression by modulating the neovasculature in ovarian TME [98]. Ovarian Cancer-Associated EVs In the course of ovarian cancer progression, intercellular communication defines the pace of neoplastic cell survival and expansion. EVs are released by almost all cell types in the TME and mediate the transfer of proteins, lipids, and nucleic acids (DNAs, mRNAs, miRNAs, and lncRNAs) between or within tumor and stroma. EVs can be subdivided into exosomes (30–150 nm) and microvesicles (100 nm–1 μm), depending on whether they are originated from multivesicular bodies or shed from the plasma membrane [99]. Stroma-released EVs can modulate cancer cell invasion and metastasis, while cancer cells also generate EVs to induce functional transition of nearby stromal cells to favor disease progression. EV

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MULTIFACETED FUNCTIONS OF THE OVARIAN CANCER MICROENVIRONMENT IN DISEASE PROGRESSION Gene symbol

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MMP1 WNT16B SFRP2 MMP12 SPINK1 MMP10 ENPP5 EREG BMP6 ANGPTL4 CSGALNACT CCL26 AREG ANGPT1 CCK THBD CXCL14 NOV GAL NPPC FAM150B CST1 GDNF MUCL1 NPTX2 TMEM155 EDN1 PSG9 ADAMTS3 CD24 PPBP CXCL3 MMP3 CST2 PSG8 PCOLCE2 PSG7 TNFSF15 C17orf67 CALCA FGF18 IL8 BMP2 MATN3 TFPI SERPINI1 TNFRSF25 IL23A

76.1 33.7 24.7 23.9 22.8 21.6 17.3 15.5 15.0 14.0 11.7 10.6 10.1 9.8 9.3 8.7 8.5 8.0 8.0 7.7 7.3 6.7 6.6 6.6 6.4 6.4 5.8 5.6 5.6 5.4 5.2 5.0 4.6 4.5 4.5 4.5 4.4 4.4 4.3 4.1 4.1 3.8 3.8 3.8 3.8 3.8 3.6 3.5

H2O2 BLEO RAD

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Apoptotic cancer cell (DDR-responsive)

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DNA-damaged stromal cell

Significantly secreted factors Cancer cell gain of functions

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FIG. 5

WNT16B and SFRP2 are significantly produced in human stromal cells during chemotherapy or radiation and promote therapeutic resistance to surviving cancer cells. (A) Genome-wide expression pattern of primary normal human stromal cells. Heatmap depicts the relative mRNA abundance after exposure of cells to typical DNA damaging agents (H2O2, hydrogen peroxide; Bleo, bleomycin; Rad, ionizing radiation). (B) Working model for cancer cell nonautonomous therapeutic resistance acquired from the TME upon anticancer treatments particularly genotoxic chemotherapy and radiation. Therapeutic agents cause apoptosis in subsets of cancer cells by eliciting a DDR, while cancer cells with DDR deficiency (DDRinsensitive, or DDR-) escape from cytotoxic attack. Simultaneously, senescence is induced in stromal cells adjacent to epithelial cells surrounding the gland, with a secretory phenotype DDSP developed after DDR events. A persistently activated signaling network is triggered by the DNA strand breaks. The DDSP is usually characterized by a spectrum of autocrine- and paracrine-acting proteins. The soluble factors reinforce the senescent phenotype in damaged cells, enhance cancer cell repopulation, with increased occurrence of tumor relapse and distant metastasis. As exemplified by the recently reported WNT16B, a handful of co-synthesized factors including SFRP2 holds the potential to serve as both a serum biomarker to determine treatment index, and a therapeutic target to minimize the TME-conferred therapeutic resistance. DDR, DNA damage response; ECM, extracellular matrix; TME, tumor microenvironment. Color images of (A) modified from Sun Y, Campisi J, Higano C, Beer TM, Porter P, Coleman I, et al. Treatment-induced damage to the tumor microenvironment promotes prostate cancer therapy resistance through WNT16B. Nat. Med. 2012;18 (9):1359–1368, with permission from Nature Medicine, copyright 2012.

formation and release are regulated by multiple factors, since either endogenous or exogenous factors can change the number, content, and type of EVs, thereby substantially altering their activities. For instance, biogenesis and release of cancer exosomes are regulated by an endosomal sorting complex required for transport, intracellular calcium levels, and structural scaffolding and subject to stimulation by stresses such as microbial attacks [100]. Signal transduction via EVs adds another level of complexity to the cell communication network as EVs simultaneously release multiple molecules impinging on signaling pathways in the recipient cell. As a subtype of noncoding RNAs, miRNAs control expression of target genes posttranscriptionally, while dysregulation of miRNAs is involved in multiple steps

of ovarian cancer development. Exosomal miRNAs isolated from the serum or ascites have potential clinical values for ovarian cancer diagnosis, prognosis, and therapeutics [101]. A former study reported the positivity of 218 out of 467 mature miRNAs isolated from ovarian cancer cells and exosomes of the same patients [102]. Among them, eight specific miRNAs had similar levels between cellular and exosomal miRNAs and exhibited correlations in a range of 0.71–0.90. Although EpCAM-positive exosomes were present in both patients with benign ovarian disease and those harboring ovarian cancer, exosomal miRNA from ovarian cancer patients displayed similar profiles and were considerably distinct from those observed in benign cases. Thus, miRNA profiling of circulating cancer exosomes can be potentially employed

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as surrogate diagnostic markers for biopsy profiling [102]. The first study investigating the miRNAs in ovarian cancer tissues and cell lines found that miR-200a, miR141, miR-200c, and miR-200b were most significantly overexpressed, while miR-199a, miR-140, miR-145, and miR-125b1 were the most downregulated miRNAs [103]. In addition, expression of miR-21, miR-203, and miR-205, whose levels were upregulated in ovarian cancer, was significantly elevated in OVCAR3 cells upon treatment by 5-aza-20 -deoxycytidine demethylating, suggesting DNA hypomethylation as a potential mechanism for miRNA overexpression [103]. Particularly, the members of miR-200 family maintain epithelial cell integrity by suppression of EMT via direct inhibition of mesenchymal transcription factors zinc finger E-box-binding homeobox 1/2 (ZEB1/ZEB2) and TGF-β, a potent inducer of EMT. Although downregulation of miR-200s in cancer cells promotes EMT and cancer metastasis, these molecules are highly expressed in ovarian cancer and cause metastasis primarily by enhancing cancer cell dissemination within the pelvic cavity [104]. Increasing lines of evidence suggest that EVs in ovarian cancer ascites are associated with immune suppression, invasion, and treatment resistance [105]. Ovarian ascites-released EVs reduce the cytotoxicity of peripheral lymphocytes and promote apoptosis of dendritic cells and lymphocytes. Further, EV-associated FAS-L purified from patient ascites triggers FAS-induced apoptosis in a T cell line, highlighting the possibility of elimination of FAS-bearing immune cells including T cells by vesicledelivered FAS-L, a process that allows immune evasion to support cancer cell survival [106]. EVs from ovarian ascites can also inhibit T cell activation by blocking the T cell signaling cascade, an effect contributed by EV-associated phosphatidylserine [107]. Depletion of HSP70+ EVs and blockade of HSP70/TLR2 interaction with A8, a specific peptide aptamer, can prevent the activation of myeloid-derived suppressor cells and abolish cancer progression in experimental mice, suggesting the potential of targeting EVs as a novel immunotherapy for ovarian cancer treatment [108]. Ovarian cancer-derived EVs carry molecules that directly regulate cancer cell migration, including CD24, EpCAM, and the soluble activated leukocyte cell adhesion molecule and soluble L1 [109–111]. Malignant ascites-released membrane vesicles contain diverse activated proteases including MMP2, MMP9, urokinase plasminogen activator (uPA), and a disintegrin and metallopeptidase domain 17 (ADAM17/TACE), together promoting ECM degradation and enhancing cancer cell invasiveness and metastasis [111, 112]. However, EVs may also carry miRNAs that positively or negatively regulate invasion and migration behaviors of cancer cells. For example, miR-6126 is released in a large amount via exosomes from both chemosensitive and chemoresistant ovarian cancer cells, and is correlated with longer

overall survival in patients with HGSC [113]. Functionally targeting integrin β1, a key regulator of cancer cell metastasis, miR-6126 acted as a tumor suppressor, and delivery of miR-6126 mimic to ECs generates decreased ovarian cancer cell invasion and migration in vitro as well as reduced tumor growth in vivo [113]. Beyond above pathological effects, EV-associated miRNAs can remarkably change the response of recipient cells to various anticancer agents. EV-mediated miR21 transfer from stromal cells including omental CAAs and CAFs to ovarian cancer cells induces resistance to paclitaxel-based chemotherapy by directly targeting the mRNA of apoptotic peptidase activating factor 1, implying inhibition of the stromal-derived miR-21 transfer is an exploitable modality in management of metastatic and recurrent ovarian cancer patients [114]. A recent study revealed that acquired SMAD4 mutations increase the chemoresistance capacity of EOCs via a novel mechanism through which platinum-resistant cell-derived SMAD4+ exosomes perpetuate an EMT phenotype and promote the expansion of platinum-refractory cell subpopulations [115]. Stable miR-433 expression in A2780 ovarian cancer cells induces a cellular senescence phenotype characterized by typical morphological changes, phosphorylated retinoblastoma (p-Rb) downregulation, and β-galactosidase activity enhancement [116]. Further, miR-433-expressing cancer cells release miR-433 into the growth media via exosomes which in turn can generate a senescence bystander effect, thus promoting the potential of recipient cells to survive chemotherapeutic treatment [116]. Exosomes released by a cisplatinresistant derivative of A2780, the CP70 cell line, were capable of driving resistance in treatment-naïve A2780 cells [117]. Similarly, cisplatin-resistant SKOV3 cells can release increased numbers of exosomes, which contain detectable annexin A3, a molecule correlated with platinum resistance of ovarian cancer cells [118].

CONCLUDING REMARKS AND FUTURE DIRECTIONS Decades of scientific and clinical efforts in ovarian cancer research provide an important baseline for the development of TME-oriented therapeutic agents that are mainly designed to interfere with tumor-stroma interactions in ovarian cancer patients. To date, many agents have achieved pronounced efficacy in improving patient survival when incorporated into traditional cytotoxic chemotherapies. However, like most human solid malignancies, ovarian cancer exhibits distinct environmental landscapes and display substantial heterogeneity [105] (Fig. 6). Even standardized treatments elicit a varying degree of responses among patient subgroups [62]. To circumvent the difficulties, emerging new techniques provide substantial benefits for translational research. For

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EVs

Metabolites

FIG. 6

Schematic illustration of a typical microenvironment that plays key roles in ovarian cancer progression. In most cases, the tumor mass is composed of single neoplastic cells and expanding cancer cell populations. Surrounding the tumor is the stroma that consists of various benign cell types, including but not limited to CAFs, MSCs, CAAs, multiple immune cell subsets, mesothelial cells, and ECs. Of note, both the tumor and stroma produce a large amount of secreted protein factors, metabolites, and EVs into the extracellular space, each substantially influencing pathological progression and holding the potential to alter therapeutic indexes in ovarian cancer clinics. CAA, cancer-associated adipocyte; CAF, cancerassociated fibroblast; EVs, extracellular vesicles; MSC, mesenchymal stem cell.

instance, engineered nanochips such as tantalum oxide nanodot arrays of 10–200 nm are now designed as artificial microenvironments, with the experimental setup and methodology specific for appraisal of ovarian cancer invasiveness [119]. This technical advancement presents an optimal diagnostic platform to investigate cancer cell behavior and to facilitate “markerless monitoring” of ovarian cancer progressiveness as a pilot model in the interdisciplinary fields of biomedical engineering and cancer research. There has been a surge in clinical trials with drugs that specifically control transmembrane or cytoplasmic enzymes, induce apoptosis and suppress angiogenesis in site-specific ovarian cancer cells, hold considerable promise to design more efficacious protocols. Future exploration in this field should focus on how to improve the practical accuracy of current therapies, to identify an increasing number of ovary-specific molecular targets for optimal effectiveness, and to develop an omics-based system for patient stratification, a highly desirable platform that underlies the personalized medicine blooming worldwide. Altogether, conceptual and technical advancement will allow physicians to make informed decisions in ovarian cancer clinics toward a thorough cure of this gynecological malignancy.

Acknowledgments We are grateful to Dr. Christian Klausen for obtaining permissions from publishers to reproduce the figures relevant for this chapter. We

want to thank the former members of Auersperg lab and current members of the Sun lab for inspiring discussion and years of experimental inputs that contribute to the scientific progress. This work was supported by grants from theNational Cancer Institute of Canada, National Key Research and Development Program of China (2016YFC1302400), National Natural Science Foundation of China (81472709, 31671425), Chinese Academy of Sciences, the National 1000 Young Talents Research Program of China, and the U.S. Department of Defense (DoD) Prostate Cancer Research Program (PCRP) (Idea Development Award PC111703) to YS.

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33 Phenotypic Plasticity and the Origins and Progression of Ovarian Cancer Lauren E. Carter, David P. Cook, Barbara C. Vanderhyden INTRODUCTION Cell plasticity—the ability for a cell to reversibly assume different phenotypes—is a common theme occurring throughout development, wound repair, and in cancer metastasis. It is often observed as an epithelial-tomesenchymal transition (EMT), allowing an epithelial cell to acquire mesenchymal traits such as migration and invasion. This cell can then reassume its epithelial phenotype through a mesenchymal-to-epithelial transition, reacquiring epithelial characteristics such as stable cell-to-cell junctions to maintain the epithelial barrier. Previously, this transition was considered a binary process where a cell could be either epithelial or mesenchymal; however, recently, this transition has been thought of as a fluid transition, where cells can lie anywhere in the EMT spectrum and express combinations of epithelial and mesenchymal traits. The concept of stemness has now been affiliated with the EMT, where cells exhibiting mesenchymal characteristics often display stem cell traits, such as the ability to form spheres in vitro. Although it has become increasingly apparent that the mesenchymal state coincides with stemness, a cell committing too far into the mesenchymal state loses these stemness traits, suggesting there is a window along the EMT spectrum that coincides with optimal stemness properties. The ovarian surface and fallopian tube epithelia are two normal tissues displaying this cellular plasticity and have the capacity to gain stem cell characteristics when pushed toward the mesenchymal state. This chapter explores the phenotypic plasticity of these two epithelia and its importance for maintaining tissue homeostasis during ovulatory wound repair. The potential consequences of this plasticity are also addressed, as both of these epithelia are putative origin sites for ovarian cancer.

The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00033-9

Plasticity may also reprogram the environment to promote cancer initiation and progression. Ovarian cancer is a heterogeneous population of cells also displaying this cellular plasticity. We discuss the contribution of plasticity to ovarian cancer progression in the context of metastasis, chemoresistance, cancer stem cells (CSCs), as well as immunosuppression. Cellular plasticity is necessary for many biological functions; however, it can also severely impact the efficacy of a cancer therapeutic. This chapter also discusses the potential of targeting this plasticity to complement current treatment strategies.

PHENOTYPIC PLASTICITY IN OVARIAN EPITHELIAL CELLS The ovarian surface epithelium (OSE) is a poorly differentiated simple epithelium surrounding the ovary. It is composed of flat-to-cuboidal epithelial cells derived from the coelomic epithelium during embryonic development [1]. Initially, the OSE was not thought to play a role in ovarian physiology so remained understudied until the late 1980s to early 2000s when studies emerged implicating the OSE as a tissue of origin for ovarian cancer [1]. Since then, it has become well understood that the OSE is a layer of cells exhibiting both epithelial and mesenchymal characteristics, and the cells composing the OSE layer have the capacity to interchange between each phenotype in response to environmental cues [1,2]. The fallopian tube epithelium (FTE) is another accepted origin of ovarian cancer and is a layer of pseudostratified epithelial cells derived from the M€ ullerian duct [3]. Unlike the OSE, the FTE represents a differentiated epithelium composed of ciliated and secretory cells. The ratio of ciliated and secretory cells is known to shift from ciliated cell

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33. PHENOTYPIC PLASTICITY AND THE ORIGINS AND PROGRESSION OF OVARIAN CANCER

dominance in the infundibulum to a secretory cell dominance in the isthmus [3]. This pattern of ciliated and secretory cells in the fallopian tube is not altered during the menstrual cycle, suggesting these cells comprise a more stable differentiated epithelium in contrast to the poorly differentiated OSE cells [3,4]. Ciliated FTE cells are not supported in cell culture conditions and over time result in a secretory FTE cell culture. Whether this is due to ciliated cells undergoing apoptosis or differentiating into secretory cells is unknown [5]. Taken together, the OSE represents a population of cells with the capacity to interchange between both epithelial and mesenchymal characteristics, whereas the FTE displays a more committed epithelial phenotype. The ability to assume reversibly different cellular phenotypes has widely been studied in the context of an EMT, and the reverse mesenchymal-to-epithelial transition, during embryonic development, cancer metastasis, and wound repair. The epithelial state maintains tissue borders and acts as an epithelial barrier and the mesenchymal state allows for cellular migration and invasion for growth or repair of tissues [6–8]. During embryonic development, epithelial cells transiently acquire a mesenchymal phenotype to allow for cellular migration. Once the cells have reached their destination, they revert to their previous epithelial state. This has classically been studied in gastrulation, neural crest cell migration, and heart morphogenesis [8]. In the context of cancer metastasis, epithelial cancer cells assume a mesenchymal phenotype to allow for migration and invasion into neighboring tissues where they revert back to their epithelial phenotype to establish a metastatic site [9]. This epithelial-mesenchymal plasticity is also observed in wound repair. In the repair of an epithelial layer, cells assume a mesenchymal phenotype surrounding the injured site to allow for migration and secretion of new extracellular matrix proteins to close the wound and mediate a process referred to as reepithelialization [10]. The OSE layer also displays this plasticity where cells exhibiting an epithelial phenotype help maintain OSE structure and during ovulatory wound repair, these cells acquire a mesenchymal phenotype to repair the wound [1,2].

THE EPITHELIAL-TO-MESENCHYMAL TRANSITION The EMT is the process of an epithelial cell acquiring the phenotype of a mesenchymal cell. This process was first described in the primitive streak of chick embryos by Elizabeth Hay in 1995 and defined as an epitheliomesenchymal transformation [11]. This event was later reclassified as a transition to emphasize the reversibility of this process and to differentiate it from cell

transformation, a process related to cancer initiation. During the EMT process, epithelial cells lose their apical-basal polarity and acquire a front-rear polarity [7,8,12]. Epithelial cell-cell junctions are lost and cell cytoskeletal reorganization occurs, resulting in the acquisition of a mesenchymal cell morphology [7,8,12]. Functionally, cells undergoing an EMT have enhanced migration and invasion and show resistance to apoptosis [8,12]. Gene expression changes commonly associated with an EMT include loss of CDH1 (E-cadherin), CLDN (Claudins), OCLN (Occludins), DSP (Desmoplakin), and PKP1 (Plakophilin) genes, all leading to a loss in epithelial barrier function [7]. Mesenchymal gene expression commonly gained during an EMT include CDH2 (Ncadherin), NCAM (Neural Cell Adhesion Molecule 1), and VIM (Vimentin), although specific genes can vary based on tissue type [7]. There have been several transcription factors (TFs) identified as master EMT-driving proteins such as the SNAIL family, basic helix-loop-helix, and ZEB family of TFs (ex: SNAI1, TWIST1, ZEB1, respectively) [7,8]. These traditional TFs are tightly regulated by mechanisms that have been evolutionarily conserved, highlighting their importance in normal cellular processes [13]. Additional TFs have been found to work in conjunction with the master EMT TFs such as Forkhead Box (FOX), GATA, and SRY Box (SOX) TF families [7]. Additional drivers of EMT have been studied, such as alternatively spliced mRNA of genes such as Cluster of Differentiation (CD) 332 and p120-catenin and noncoding RNAs (microRNAs and long noncoding RNAs) such as miR-34 and H19, respectively [14–17]. With the emergence of high-throughput techniques, such as RNA-Seq and ChIP-Seq, more complex regulation of the EMT process is being uncovered and increasingly more interactions being identified as regulators of this program. Recently, many studies have demonstrated that the EMT is best represented by a spectrum with the epithelial phenotype at one end, the mesenchymal phenotype at the other end, and a range of intermediate, or partial EMT in between [6–8,18]. In the intermediary states, also referred to as “metastable” states, cells can possess varying levels of both epithelial and mesenchymal traits [6]. Huang et al. used 43 ovarian cancer cell lines to demonstrate the range of EMT states [19]. Using EMT-related gene expression, they classified these tumor cell lines into 4 categories: epithelial, intermediate epithelial, intermediate mesenchymal, and mesenchymal represented by 20.9%, 41.9%, 18.6%, and 18.6% of cell lines, respectively [19]. They found that different EMT-TFs peaked in expression levels in these categories, suggesting these EMT-TFs have different weights in dictating where the cell line falls within the EMT spectrum [19]. The next year, Tan et al. developed an EMT scoring method using gene expression profiles of a variety of cancer cell types to estimate quantitatively where a particular tumor or cell line falls

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THE EPITHELIAL-TO-MESENCHYMAL TRANSITION AND STEMNESS

within this EMT spectrum [20]. Most recently, Jolly et al. introduced the concept that the EMT spectrum is multidimensional and different EMT modulators contribute to different EMT characteristics on this spectrum. For example, the EMT characteristics migration and invasion can be represented as different branches on the EMT spectrum and are regulated by different sets of EMT-TFs [21]. Taken together, it is becoming increasingly apparent that the EMT is a fluid state regulated by numerous factors, and cells have the capacity to shift their positioning on the spectrum based on their environmental cues (Fig. 1A).

formation, which are two characteristics of stem cells [22]. Guo et al. further characterized this phenomenon using transient ectopic expression of Slug (another EMT-TF), showing these stem cells were functional in vivo in a mammary gland reconstitution assay [23]. This efficiency was enhanced when another EMT-TF commonly associated with stemness, Sox9, was also expressed [23]. This study suggests that regulators of EMT can drive this stemness phenotype synergistically [23]. In 2015, Schmidt et al. showed that constitutive Twist1 expression induced an EMT in mammary epithelial cells; however, transient expression was required to promote stemness [25]. The authors observed that removal of Twist1 did not fully restore cells to an epithelial state (mesenchymal-to-epithelial transition, MET), but maintained cells in a partial EMT state where they were “primed” for stemness, suggesting there is a window of stemness within the EMT spectrum [25]. This concept has also been inferred by Jolly et al. using a theoretical modeling of an EMT where both limits of the EMT spectrum have cells exhibiting less stemness than the intermediate zones [26]. The authors further elaborated upon this idea by showing that the positioning of the window of stemness on the EMT spectrum is not universal for all cell types and this positioning can be set using EMT driving factors, but “fine-tuned” using

THE EPITHELIAL-TO-MESENCHYMAL TRANSITION AND STEMNESS To further the EMT spectrum complexity, it has been demonstrated that cells undergoing an EMT acquire stem cell characteristics (stemness) [22–25]. Mani et al. were the first to publish this finding in 2008 using nontransformed cells [22]. Immortalized human mammary epithelial cells induced to undergo an EMT via Transforming Growth Factor Beta-1 (TGFB1) treatment or using ectopic expression of TWIST or SNAIL were found to acquire stem cell marker expression and increased mammosphere Epithelial

Intermediate phenotypes

Tight cell-cell contact Apical-basal polarity Basement membrane anchorage

EMT

Mesenchymal

Loss of polarity and cell-cell contacts Cytoskeletal rearrangement Enhanced migration Enhanced stemness characteristics

MET

(A) Decoupling morphology and stemness effects (”window of stemness”)

Cell type #1

Stemness

Stemness

Strict correlation between morphology and stemness

Cell type #2

Cell type #3 Epi

Morphology

Mes

Epi

Morphology

Mes

(B) FIG. 1 (A) A schematic of the epithelial-mesenchymal transition, highlighting a continuous transition from an epithelial to a mesenchymal state, along with characteristics associated with each state. (B) Two models of the relationship between stemness and morphological characteristics associated with an EMT. Historically, these traits have been thought to correlate quite strongly (left model); however, recent work suggests that stemness is often optimal in an intermediate state, which may be context-dependent (“window of stemness” model; right). EMT, epithelial-to-mesenchymal transition; MET, mesenchymal-to-epithelial transition.

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modulating factors such as OVOL. OVOL couples with traditional EMT-driving factors to inhibit a full mesenchymal phenotype but simultaneously enhances the stemness phenotype [27]. This concept of a stemness window within the EMT spectrum has been shown in ovarian cancer cell lines where those in an intermediate EMT state possess enhanced spheroid formation and are anoikis resistant, both stemness characteristics [19]. Taken together, it appears that within the EMT spectrum there is a window of stemness whose specific location is modulated by EMT-driving factors that work together to regulate where this window lies (Fig. 1B).

STEMNESS IN THE OVARIAN SURFACE EPITHELIUM Identification and Characterization of Stemness OSE cells are thought to contain a stem cell population that is present to maintain tissue homeostasis [28]. Since 2008, several studies have evaluated common stem cell characteristics to identify this population of cells. DNA label retention has been used to identify slow-cycling somatic stem cells that exhibit asymmetric division and retain a label for long periods of time. In comparison, lineage-committed daughter cells (which proliferate more rapidly) dilute out the label as they proliferate. Szotek et al. were the first to report a population of stem-like mouse OSE cells using this technique [29]. Side population analysis is another common technique used to identify a population of stem cells and can be identified using the ATP-binding cassette (ABC) pump inhibitor, Verapamil. ABC pumps are abundant in stem cells and are therefore able to efflux dyes such as Hoechst dye efficiently, whereas differentiated cells have a limited ability to do so. Szotek et al. found their population of label-retaining cells was enriched when isolating the side population present in mouse OSE cells [29]. The following year, Bowen et al. published microarray and immunohistochemical data of human OSE cells, showing activation of stem cell quiescence and regulation pathways as well as expression of stem cell markers within these cells, suggesting these putative stem cells exist in both mouse and human ovaries [30]. From 2012 to 2014, several publications further characterized this stem cell population in both mouse and human using in vitro assays such as sphere formation, and in vivo assays such as lineage tracing and label retention [31–35]. There have even been reports of OSE stem-like cells with the capacity to differentiate into additional cell types, including oocyte-like structures [36–41]. Although many studies have identified stem cell characteristics in the OSE, there has yet to be a consensus on which marker(s) accurately represent this population of cells. One possible reason for this

discrepancy in stemness markers, may be that the OSE does not have a fixed stem cell population but a transient one, that can be induced or expanded based on environmental cues.

Regulation of OSE Stemness Isolating OSE stem cells based on their gene expression may enrich for stem cell characteristics; however, there has not been any demonstration that combining stem cell marker expression purifies this population of cells, like it does in the hematopoietic system—a hierarchical stem cell model. This suggests the OSE stem cell population is best represented as a state rather than a defined population of cells, where these cells have the capacity to interchange between a “less differentiated state” exhibiting stem cell characteristics and a “more differentiated” cell state exhibiting more committed epithelial cell characteristics. Gamwell et al. showed that culturing mouse OSE cells negative for a stem cell marker Stem Cell Antigen 1 (Sca1/Ly6a) revert back to a Sca1-positive state, a marker of cells exhibiting stem cell characteristics, exemplifying the plasticity of this cell state [31]. Furthermore, the authors found that treating mouse OSE cells with TGFB1 (a known EMT inducer) increases stem cell characteristics, such as sphere formation and stem cell marker expression in vitro, indicating this cell state is regulated by factors such as those driving an EMT. Studies examining EMT regulation and the stemness phenotype in OSE are summarized in Tables 1 and 2. To this date, there have not been any direct studies in the OSE relating EMT and stemness, but this relationship has been established in other tissues as described above. As there is no agreement or coordination of stemness markers in the OSE, but there are clear stem cell phenotypes, perhaps the window of stemness is best represented by adding a second dimension to the EMT spectrum, where cells lie within the EMT spectrum based on their epithelial/mesenchymal genes, and are shifted into the stemness dimension based on their stemness gene expression. This theory would allow for cells to exhibit similar functional readouts, while expressing different genes and would explain the discrepancy in some experimental findings (Fig. 1B).

STEMNESS IN THE FALLOPIAN TUBE EPITHELIUM Like the OSE layer, the FTE is also thought to contain a stem cell population that is present to maintain tissue homeostasis, although less is known about this population of cells. Wang et al. identified a population of label

VI. HUMAN OVARIAN CANCER

TABLE 1

Overview of EMT/MET Regulation in OSE and FTE Cells

Cell type

In vitro/ in vivo

Species

Treatment

EMT/MET characteristics

Ref.

OSE

In vitro

Human [primary culture]

Epidermal growth factor [EGF]

Mesenchymal morphology " Motility " Secretion of promatrix metalloproteinases " ERK, ILK, AKT, and GSK-3 signaling

[142]

OSE

In vitro

Human [immortalized cells]

Exogenous E-cadherin expression

Epithelial morphology " Adherens and tight junctions " Keratin expression

[143]

OSE

In vitro

Human [primary culture]

EGF + hydrocortisone

Mesenchymal morphology # Keratin expression " Collagenous extracellular matrix

[144]

OSE

In vitro

Human [primary culture]

Bone Morphogenetic Protein 4

No change in cell motility

[145]

Noggin OSE

In vitro

Human [primary culture]

TGFB1

Prevented formation of epithelial barrier # E-cadherin, Claudin 1, Occludin, Crumbs 3 " Snail, N-cadherin, Slug

[146]

OSE

In vitro

Mouse [primary culture]

Exogenous Pax8 expression

Mesenchymal morphology " Migration " N-cadherin and Fibronectin

[147]

OSE

In vitro

Human [primary culture]

Collagen gels

Mesenchymal morphology

[148,149]

FTE

In vitro

Mouse [primary culture]

Pax8 knockdown

No changes in migration or apoptosis

[147]

FTE

In vitro

Mouse [primary culture]

TGFB1

Mesenchymal morphology " Migration " Snail and # E-cadherin

[45]

EMT, epithelial-to-mesenchymal transition; FTE, fallopian tube epithelium; MET, mesenchymal-to-epithelial transition; OSE, ovarian surface epithelium; TGFB1, Transforming Growth Factor Beta-1.

TABLE 2

Overview of the Regulation of OSE and FTE Stemness

Cell type

In vitro/ in vivo

OSE

OSE

Species

Treatment

Stemness characteristics

Ref.

In vitro

Human [primary culture]

Follicular fluid

Formation of primitive oocyte-like cells positive for alkaline phosphatase and markers [150] of pluripotency [SOX2, SSEA4, OCT4A, NANOG, NANOS, STELLA, CD9, LIN28, KLF4, GDF3, and MYC]

In vitro

Mouse [primary culture]

Follicular fluid

" Stem cell marker expression [Sca1]

TGFB1

# Proliferation, " sphere formation, " Sca1 expression, converts SCA1 to SCA1+ cells

LIF

" Sphere formation, # Sca1 expression

[31]

OSE

In vivo

Mouse

Ovulation

Label retaining cells [stem cells] are proliferative postovulation

[29]

OSE

In vivo

Mouse

Ovulation

Ovulating OSE regions exhibit mitotically active LGR5+ cells [stem cells] LGR5+ cells respond to local Wnt signals

[33]

OSE

In vivo

Mouse

miR-34 family and miR-376b

# in the stem cell population [ALDH+ cells]

[32]

OSE

In vitro

Mouse [primary culture]

Exogenous Pax2 expression

# Sphere formation and stem cell marker expression [CD44, Sca1, Lgr5]

[45]

FTE

In vitro

Mouse [primary culture]

Serum

Label retaining cells " expression of endometrial, proximal, and distal oviductal specific [42] genes

FTE

In vitro

Mouse [primary culture]

TGFB1

" Stem cell marker [CD44 expression]

Pax2 knockdown " Stem cell markers [CD44, Sca1, CD133] " sphere formation

FTE, fallopian tube epithelium; OSE, ovarian surface epithelium; TGFB1, Transforming Growth Factor Beta-1.

[45]

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33. PHENOTYPIC PLASTICITY AND THE ORIGINS AND PROGRESSION OF OVARIAN CANCER Auersperg et al. [34] Human stem cell markers (NANOG, SFRP1, LHX9, ALDH1A1, and ALDH1A2) detectable in human OSE in vivo.

Bowen et al. [30]

Ovarian surface epithelium

Detection of adult stem cell-associated gene expression in human OSE.

Flesken-Nikitin et al. [32] ALDH1+ population of cells enriched at the murine ovarian hilum with enhanced sphere forming ability. Cells proliferate in response to estrus cycling.

Szotek et al. [29] Label-retaining cells that exhibit enhanced colony formation. Cells enriched in a cytoprotective side population.

Gamwell et al. [31] SCA1+ mouse OSE exhibit enhanced sphere formation and are expanded by factors in follicular fluid (TGFB1, LIF).

LGR5+ cells located around ovulatory wound sites in the mouse ovary that proliferate in response to estrus cycling and contribute to OSE homeostasis. LGR5+ expression confirmed in OSE of human ovaries.

Ng et al. [33]

Wang et al. [42] Label-retaining cells in the distal oviduct exhibit enhanced sphere formation and differentiation upon serum stimulation in vitro.

Paik et al. [43]

Fallopian tube epithelium

Ng et al. [33]

Human FTE cells lacking markers of differentiation in the basal cells of the distal fallopian tube. Express CD44 and have enhanced sphere formation in vitro.

Patterson and Pru [44]

Rare KRT8+ cells expressing LGR5 at base of mouse oviductal epithelium. LGR5 expression confirmed in human FTE.

Alwosaibai et al. [45] Mouse OSE express the stem cell marker CD44 in vitro. Expression is repressed by the transcription factor PAX2.

Alwosaibai et al. [45] Oviductal epithelial cells express the stem cell marker CD44, whose expression is increased by TGFB1. PAX2 deletion increases marker expression and sphere forming efficiency.

Auersperg et al. [34] Human stem cell markers (NANOG, SFRP1, LHX9, ALDH1A1, and ALDH1A2) detectable in human FTE in vivo.

Label-retaining FTE cells in the distal oviduct that lack markers of differentiation and express the stem cell marker KIT.

FIG. 2 A timeline of publications describing putative stem cell populations of the ovarian and fallopian tube/oviductal epithelia. The field experienced a burst of activity between 2012 and 2014; however, this has been followed by several years of inactivity.

retaining cells located in the epithelium of the distal end of the mouse fallopian tube (oviduct) [42]. These cells formed spheres in vitro and were able to differentiate into glandular structures expressing mature Mullerian epithelial cell markers [42]. The ability of these cells to contribute to FTE homeostasis was not assessed, but these cells were able to be differentiated into cell lineages of the endometrium, proximal and distal oviduct [42]. Paik et al. identified a population of stem-like cells in the distal end of the human fallopian tube expressing stem cell markers and lacking expression of ciliated and secretory cell markers [43]. These cells also had sphere forming potential, suggesting this population of stem-like cells exists in the human fallopian tube as well [43]. Patterson and Pru also demonstrated label retention in the distal end of the mouse oviductal epithelium [44]. Additionally, there has been a population of cells exhibiting stem cell characteristics located at the hilum region of the mouse ovary, where the OSE and oviductal epithelium meet, that is, slow cycling expresses stem cell markers and is capable of sphere formation [32]. Additional stem cell markers have been found in the distal end of the mouse oviductal epithelium, such as Lgr5 [33]; however, further characterization of these cells and their regulation has not been assessed. Recently, a study showed that the stem cell characteristics in the oviductal epithelium can be enhanced with Pax2 knockdown or TGFB1 treatment, which works in part by decreasing Pax2 expression [45]. This study directly implicates an EMT with the acquisition of stemness in the oviductal epithelium.

Overviews of the characterization of EMT and stemness in the FTE are presented in Tables 1 and 2. These studies show that both OSE and FTE interchange between epithelial and mesenchymal phenotypes and, when in a more mesenchymal state, exhibit stemness characteristics. Constructing a timeline of discoveries on OSE and FTE stemness emphasizes a surge in studies between 2012 and 2014 exploring the putative stem cell population in both of these epithelia. Despite this, however, momentum was lost as no further studies have been published since, with the exception of Alwosaibai et al., who found that knockdown of the transcription factor Pax2 in oviductal epithelial cells increased stem cell characteristics, and forced expression in OSE cells reduced the features of stemness [45] (Fig. 2). This lull in activity since 2014 may be random, or perhaps it reflects the time taken to collect experimental findings. It may also reflect the recent shift the field has taken to focus on the FTE as the origin of ovarian cancer, and since in vitro culturing methods of FTE ciliated cells have not been optimized, progress is slowed.

PHENOTYPIC PLASTICITY IN OVULATORY WOUND REPAIR Ovulation, the process where the mature oocyte leaves the ovary for fertilization in the fallopian tube, is a cyclic process that is composed of three stages: the preovulatory, ovulatory, and postovulatory phases [28]. In the preovulatory phase, the dominant follicle continues to

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PHENOTYPIC PLASTICITY IN OVULATORY WOUND REPAIR

develop to the antral or Graafian stage where it is ready for ovulation. During this development, the follicle size is dramatically increased producing a stigma in the ovarian tissue surrounding the follicle. In the events leading up to ovulation, the OSE layer surrounding the stigma proliferates to accommodate this change in ovarian size, whereas the OSE cells surrounding nonovulating sites remain nonproliferative [46,47]. Ovulation is triggered by a surge in luteinizing hormone (LH). The LH surge induces OSE cells to produce proteases, such as urokinase-type plasminogen activator, which activates a proteolytic cascade that degrades extracellular matrix proteins and induces apoptosis at the apex of the ovarian stigma [48]. This epithelial cell breakdown facilitates the rupture in the OSE layer, allowing for ovulation to proceed. Once the cumulus-oocyte complex is released from the ovary, the ovulatory wound is repaired in as little as 12 h in mice [47,49].

Reepithelialization Wound repair is a complex event that requires synergy of multiple components to be successful. Reepithelialization is one step in this process where cells on the edge of the wound site migrate to reestablish the damaged epithelium [6]. After ovulation, OSE cells surrounding the wound site continue to proliferate and assume a mesenchymal phenotype allowing for the secretion of collagen and extracellular matrix proteins [48]. Cell plasticity is also important for reepithelialization of the OSE layer. OSE cells need to assume a mesenchymal morphology to facilitate the repair, however, require the ability to reestablish their epithelial phenotype once the repair is completed to form the epithelial barrier. Recently, studies in the skin have shown that a partial EMT is induced during wound repair [10]. This transient reprogramming of the wound edge cells facilitates their migration and invasion into the wound site as a cohesive cohort and not as individual cells [8]. During this transient reprogramming, intermediate filaments are retracted, breakdown of the basement membrane occurs, and epithelial cells lose their polarity. Once cell migration has taken place, these cells reform their cell-substrate contacts [50]. During this repair process, cells maintain their cytokeratin expression and desmosomes, further implying these cells undergo a partial EMT [8]. This partial EMT can be seen in the OSE cell layer, which maintains expression of keratin 8 and E-cadherin throughout ovulatory wound repair [51]. During ovulatory rupture, follicular fluid from the antral follicle bathes the OSE layer surrounding the rupture site. A recent study assessed the composition of human follicular fluid and found proteins belonging to several functional groups such as insulin growth factor

535

(IGF) and IGF-binding protein families, growth factors and related proteins, receptor signaling, defense/immunity, antiapoptotic proteins, matrix metalloproteaserelated proteins, and complement activity [52]. The authors found that once ovulation is induced (following human chorionic gonadotrophin treatment), the composition shifted to include more proteins belonging to the protease inhibition, inflammation, and cell adhesion families [52]. Many of these follicular fluid components are known inducers of an EMT such as TGFB1 and IGF. These components may help facilitate the transient reprogramming to an EMT state to aid in wound repair. OSE undergoing reepithelialization around the wound site also gain expression of the stem cell marker Lgr5, suggesting this partial EMT allows for reepithelialization and also induces stemness, necessary to maintain homeostasis [33]. Furthermore, Gamwell et al. showed that treating OSE cells with follicular fluid enhances their stem cell characteristics, suggesting ovulatory wound repair expands this population [31]. The concept that ovulation changes the OSE stem cell profile partially explains why there has not been agreement in the gene markers for this population of cells. Different factors produced in the surrounding ovarian tissue interact with the OSE layer to promote this stem-like state to maintain tissue homeostasis. Since these factors are complex and transient within the ovary, it is not probable that one marker of stemness will be found that represents all stem cells, but it is possible to describe these cells as a state within the spectrum.

Inflammation Another important player in wound repair is the immune system. In adult tissues, wound repair relies heavily on a functional immune response. Wound repair is hindered when the immune response is impaired [10]. For example, inhibiting macrophage function has been shown to delay tissue repair in rabbits and mice [53,54]. Inflammation occurring after an epithelial wound recruits primarily neutrophils and macrophages to the wound site, which work together to clear the wound debris [10]. In the ovary, these immune cells home to the forming corpus luteum after ovulation in the mouse, rat, and human [55–57]. Inflammation also supports reepithelialization through the secretion EMT-driving factors such as TGFB1 and tumor necrosis factor (TNF) alpha, which act on the epithelial cells surrounding the wound site [10,58,59]. As mentioned above, human follicular fluid also becomes more inflammatory at ovulation, suggesting follicular fluid contributes to the inflammatory response [52]. Taken together, inflammation occurring during wound repair is important to facilitate this repair by altering the ovarian niche to be more inflammatory and promote an EMT in the repairing epithelium.

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Potential Consequences of Cell Plasticity Ovulation is a cyclic event, occurring once every 28 days in women of reproductive age. It is also the primary nonhereditary risk factor for ovarian cancer, with the number of ovulatory cycles being proportional to ovarian cancer risk [60]. This suggests that ovulation, or ovulatory wound repair, can lead to ovarian cancer initiation. During each cycle, the repairing OSE layer proliferates and undergoes a partial EMT. During this time, the cells surrounding the wound site also interact with the inflammatory agents present in the follicular fluid. Cells exhibiting a mesenchymal phenotype display resistance to apoptosis and therefore can survive harsh environmental conditions; however, this cell survival may allow the accumulation of DNA damage over time. Murdoch and Martinchick assessed the surviving OSE cells surrounding the ovulatory rupture site and found 8-oxoguanine modifications in these cells, indicating DNA damage [61]. As these cells are undergoing an EMT also have enhanced stemness characteristics, it is conceivable that these cells may develop DNA mutations that are propagated over time and can lead to ovarian cancer initiation. The inflammation accompanying wound repair is tightly regulated since its dysregulation could have severe consequences. Improper wound repair in other tissues such as the liver, kidney, and lung results in tissue scarring and fibrosis, leading to chronic inflammation [6]. This chronic inflammation results in persistent secretion of EMT-driving factors and maintenance of the EMT state in the epithelium [12]. As EMT is related to stemness characteristics, chronic inflammation in the ovary would also increase the stem cell characteristics in the OSE and distal FTE, which over time could transform into a cancer initiating cell. Briley et al. reported that fibrosis in the mouse ovary increases with age and number of ovulatory cycles [62]. Although this has yet to be established in humans, ovarian fibrosis is known to be associated with reduced fertility in women with polycystic ovarian syndrome [63]. It is possible that fibrosis in the human ovary increases with age and the associated chronic inflammation could promote both an EMT and stemness in the OSE and FTE. This combination of events theoretically could provide a microenvironment conducive to cell transformation and the development of neoplastic lesions. This hypothesis could help to explain why the median age of ovarian cancer detection is postmenopausal, after the age-associated accumulation of postovulatory repair events results in ovarian fibrosis and inflammation [60]. The FTE is another site of origin for ovarian cancer, and is also affected by ovulation. The follicular fluid that is expelled from the ovary comes into contact with the distal FTE cells. Like in the OSE layer, the follicular fluid components may drive an EMT phenotype, induce DNA damage, and promote stemness in the FTE. This concept

is supported by Alwosaibai et al. who found that treating FTE cells with TGFB1, a component of follicular fluid, induced an EMT and increased the expression of stem cell markers [45]. Interestingly, these effects were mediated in part by downregulation of Pax2, a transcription factor commonly lost in serous tubal intraepithelial carcinomas, a precursor lesion for ovarian cancer. Furthermore King et al. detected proinflammatory macrophages in the oviduct of superovulated mice as well as increased levels of phospho-gH2A.X, a marker of DNA damage, in the oviductal epithelial cells [64]. Taken together, the plasticity exhibited during wound repair by both the OSE and FTE layers may enable wound repair to take place and work to maintain tissue homeostasis. Under the inflammatory conditions of wound repair, however, it may change the ovarian niche to result in more DNA damage and promote transformation.

HETEROGENEITY OF OVARIAN CANCER There are many different types of ovarian cancer, each characterized by distinct histological and molecular characteristics [65,66]. Much of the variation between these subtypes is generally accepted to be due to differences in the cancer’s cell-of-origin or specific genetic aberrations [67,68]. For example, there is evidence that endometrioid ovarian cancer is derived from endometrial tissue of atypical endometriosis, while high-grade serous carcinoma (HGSC) is thought to originate from either the FTE or OSE, and HGSC are often associated with TP53 mutations, whereas low-grade carcinomas are not [67,68]. Over the last decade, advances in large-scale genomics have enabled interrogation into the variation that exists across tumors of the same histological subtype. The Cancer Genome Atlas profiled the transcriptome of 489 HGSC tumors and noted that the tumors fell into four distinct clusters: differentiated, immunoreactive, mesenchymal, and proliferative—each named for the gene expression patterns that define the cluster [69]. A study by Tan et al. [70] further refined this classification by including gene expression data sets from over 2500 additional tumor samples to their analysis and once again found that tumors were members of distinct molecular subtypes. The expression patterns associated with each subtype resembled the clusters previously defined by The Cancer Genome Atlas, but were largely driven by the epithelial-, mesenchymal-, or stem cell-associated gene expression patterns, so they were named Epi-A, Epi-B, Mes, Stem-A, and Stem-B [70]. Importantly, both studies found that the molecular subtype of the group can impact clinical outcomes. For example, the Stem-A and Epi-B classifications were shown to be prognostic factors, independent of other factors, such as tumor stage

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CONSEQUENCES OF THE EMT ON OVARIAN CANCER PROGRESSION

[70]. Additionally, the efficacy of various treatments can be dependent on the molecular subtype: Stem-A tumors are particularly susceptible to microtubule assembly inhibitors [70], while Mes tumors are selectively susceptible to inhibition of the receptor tyrosine kinase AXL [71]. While it is evident that ovarian tumors of the same histological subtype can be quite heterogeneous, and that this contributes to tumor progression and patient prognosis, the source of this heterogeneity is not clear. All molecular profiling used to define these subtypes has relied on technologies that make measurements from bulk populations of cells (e.g., a tumor core), and as a consequence, are sensitive to differences in the cellular composition of the tumor. For example, a recent study found that the mesenchymal and immunoreactive classifications from The Cancer Genome Atlas can likely be attributed to tumors with a higher stromal content, rather than a unique expression signature from the carcinoma cells themselves [72]. However, while defining the Epi-A, Epi-B, Mes, Stem-A, and Stem-B subtypes from tumor samples, Tan et al. also found that cultured ovarian cancer cell lines, which are likely relatively pure cancer cell populations, could also be clustered into these five subtypes while maintaining a relatively high correlation with expression profiles from tumor samples of the same subtype [70]. This suggests that while the tumor microenvironment can certainly contribute to the variation between molecular profiles, the carcinoma cells themselves can also be phenotypically diverse. What is not clear is how this diversity arises. It is possible that these cells arise from a different cell-of-origin; however, there are only two putative sources of HGSC—the FTE and OSE (or inclusion cysts derived from them)—which does not account for the diversity of molecular subtypes. It could also be that tumors of the same subtype share a common mutational profile that may drive the particular phenotype, but this has yet to be shown. What seems likely is that the cancer cells are transcriptionally and epigenetically programmed by tumor microenvironment throughout their initiation and progression, driving this phenotypic diversity. As discussed previously in this chapter, normal ovarian epithelial cells are quite plastic and readily shift between epithelial and mesenchymal phenotypes to maintain tissue homeostasis, so it seems likely that ovarian carcinoma cells retain this property. Unfortunately, evidence of ovarian carcinoma cell plasticity in vivo is sparse, and much of the work to support this hypothesis has been demonstrated with ovarian cancer cell lines in vitro. However, over the last 14 years, since the earliest studies exploring the relationship between the EMT and ovarian cancer progression, there has been an accumulation of several hundred studies that explore the factors that drive this phenotypic shift.

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The key drivers of the EMT in ovarian carcinoma cells seem to be largely conserved across other normal and cancer cell types. The classic EMT-TFs SNAI1 and SLUG were the first factors shown to induce an EMT in ovarian cancer cells [73]. Their ectopic expression in a cultured HGSC cell line induced the morphological traits of an EMT, as well as enhanced their ability to migrate and invade through a matrigel substrate in vitro [73]. Soon after, TWIST1 [74], TWIST2 [75], ZEB1 [76], and ZEB2 [76,77] were each implicated to have similar roles. Many of the published studies in this field have focused on specific components of the signaling pathways that induce an EMT and have found commonalities with other epithelial tissues and tumors. Receptors and kinases of many of the major signaling pathways, including the TGFB [78], WNT [79], PI3k-AKT [80], ERK [81], and JAK/Stat pathways [81], have been shown to induce an EMT in ovarian carcinoma cells. Additionally, several microRNAs and long noncoding RNAs have been shown to modulate the expression of EMT TFs. The miRNA-200 family [76], 125a [82], 429 [83], 187 [84], 29b [85], 101 [86], 7 [87], 150 [88], 506 [89], 373 [90], 186 [91], 153 [92], 203 [93], 1181 [94], 31 [95], 30d [96], 382 [97], 340 [98], 125b [99], 137 [100], and 34a [100] each have been shown to inhibit an EMT in ovarian cancer cells, whereas only miRNA-181a [101], 23a [102], 186 [91], and 21 [103] have been shown to induce one. Many of these microRNAs have been shown to function directly through targeting the transcripts of the classic EMT-TFs. Less is known about long noncoding RNAs in ovarian cancer; however, HOTAIR has been shown to induce an EMT [104], while the noncoding HOXA11-AS prevents the transition [105]. It is interesting to appreciate the imbalance of factors that drive cells along the epithelial-mesenchymal continuum: there seem to be far more TFs and signaling molecules that actively push cells to be more mesenchymal rather than epithelial. Similarly, there seem to be more microRNAs, which are typically repressive in nature, that make cells more epithelial. Perhaps this is a bias in research (e.g., labs research EMT more than MET), or perhaps this suggests that the regulatory systems positioning cells along the epithelial-mesenchymal continuum are not two opposing gene networks (EMT vs MET programs), but rather a single, tunable gene network, and the phenotype of each cell is defined by degree of its activity.

CONSEQUENCES OF THE EMT ON OVARIAN CANCER PROGRESSION Metastasis Many of the studies identifying regulators of the EMT also assessed functional characteristics of the cells in addition to monitoring cell morphology and gene

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expression. Consistent with other EMT literature, most describe the cells’ increased ability to migrate in vitro using scratch-wound assays and increased invasion through an extracellular matrix (e.g., matrigel). It has long been thought that undergoing an EMT is critical for cancer cells to break from the primary tumor, invade through adjacent tissue, and metastasize throughout the body [106]. In fact, modulating the EMT status of ovarian carcinoma cells does seem to affect metastasis in xenograft models. For example, knockdown of SNAI1 in an ovarian cancer cell line led to a near-complete inhibition of metastasis in an orthotopic mouse model [107]. In this same model, SNAI1 expression also tended to be higher in metastatic tumors than primary ovarian tumors [107]. Recently, the EMT-metastasis hypothesis has been met with criticism, largely because findings have hinged on experimentally inducing or inhibiting the EMT. So while more-mesenchymal cells seem to have an enhanced ability to metastasize, it is not clear if carcinoma cells dynamically activate an EMT to facilitate metastasis, or to what extent the EMT-associated genes are required to allow metastasis to occur. Two recent studies published in Nature used conditional knockouts or genetic lineage tracing, fluorescently labeling cells upon the expression of EMT-TFs, to demonstrate that the cells of metastases from breast [108] and pancreatic [109] cancer mouse models had not undergone an EMT. However, these studies are not without their own limitations. The genetic manipulations in both studies were limited to a small number of EMT-associated genes. Knockout of a single EMT-TF may not be sufficient to block the metastasis component of an EMT in this model, and lineage tracing based on a single gene may not capture all cells that had undergone an EMT. However, Fischer et al. [108] also demonstrated that overexpression of miR-200, which inhibits the expression of multiple EMT genes, still failed to affect metastasis, further supporting that at least in some contexts, the EMT may be dispensable in metastasis. In ovarian cancer models, there has not been much focus on tracking cells throughout the process of metastasis, and most EMT-metastasis studies have relied on overexpressing or knocking down gene expression. Rafehi et al. did find that ovarian cancer cells, when switched from adherent culture to spheroid culture in suspension, had elevated expression of mesenchymalassociated genes, and mostly restored epithelial gene expression when placed back in adherent culture conditions [110]. The ability for the cells to grow in suspension was also reduced when the cells had been exposed to a TGFB signaling inhibitor. While this is consistent with cells undergoing an EMT when transferred to suspension culture, it is also possible that this is not a dynamic response, but rather, the process is simply selective for cells that already had a more-mesenchymal phenotype.

However, the ability to restore epithelial gene expression when the spheroids were transferred back to adherent culture is supportive of dynamic plasticity, rather than selection of static cell populations.

Chemoresistance Another cancer-associated trait modulated by the EMT is sensitivity to platinum-based chemotherapy—a prevalent issue across a variety of cancers [111]. Many of the studies that identified regulators of the EMT in ovarian cancer also implicated these factors in promoting chemoresistance. The EMT-inducing TWIST1 [112], STAT3 [113], FOXM1 [114], and more, each endow ovarian carcinoma cells with an increased resistance to cisplatin, whereas miRNA-186 [91] and KLK4 [115]—each of which induces an MET—render cells more sensitive to cisplatin. Given this, therapeutically targeting the EMT seems like a logical strategy to sensitize ovarian carcinomas to chemotherapy [116–118]. Interestingly, these findings are all confounded by one large study that performed gene expression microarrays on 46 human ovarian cancer cell lines [119]: each cell line was assigned a binary classification of “epithelial” or “mesenchymal,” and was assessed for sensitivity to cisplatin, and the gene expression changes that occurred in response to a 48-h exposure to a cell-line-specific GI50 dose [119]. They surprisingly found that epithelial-classified cell lines were less sensitive to cisplatin than the mesenchymal lines, with a higher GI50 dose on average and a reduced apoptotic response [119]. The authors mention that this finding may be the result of differences in “acquired” and “inherent” EMTs; however, these findings have yet to be consolidated with the immense body of work relating an EMT phenotype with resistance to chemotherapy.

Cancer Stem Cells Related to the chemoresistant trait is the notion of CSCs. It has been hypothesized that, like in tissue development, the cells comprising tumors are hierarchically organized, with self-renewing stem cells populating the tumor through cell division and subsequent differentiation of daughter cells [120]. A logical extension is that not all tumor cells are equally capable of forming new tumors. In fact, in many types of cancer, it has been demonstrated that cells with certain gene expression signatures are more capable of forming tumors in xenograft models than cells lacking the given signature. While limiting dilution xenograft tumor models are currently the gold standard for assessing this phenotype, the ability to form clonal, self-renewing spheroids in suspension culture is the most commonly used proxy for this trait. Additionally, resistance to chemotherapy is a feature of CSCs,

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attributed to the expression of ABC transporters, enhanced DNA damage response, and more [121]. In ovarian cancer, the expression of several marker genes, albeit in different combinations in different studies, has been attributed to putative CSCs, including CD44, CD24, CD133, ALDH1, CD117, and EPCAM [122]. In the first study that demonstrated that the EMT endows cells with stem cell characteristics, Mani et al. proposed that if the EMT provides normal epithelial cells with stem cell characteristics, perhaps it also provides carcinoma cells with these traits [22]. The authors demonstrated that in limiting dilutions, transformed mammary epithelial cells with ectopic expression of the EMT factors Snail or Twist formed more tumors than cells that had not undergone an EMT [22]. Since this report, it has been demonstrated that the EMT promotes cancer stem cell characteristics in various cancers, including pancreatic [123], colon [124], hepatocellular [125], lung [126,127], prostate [128], and head and neck [129]. While there are quite a few studies about putative ovarian CSCs and their characteristics, less is known about their relationship with the EMT in ovarian cancer. There are a few pieces of evidence to support that the EMT may actively contribute to the CSC phenotype. The EMT-inhibiting miRNA-200 has been shown to reduce the tumorigenicity of a population of putative ovarian CSC [130]. Additionally, CD24-expressing ovarian CSCs are more invasive and express higher levels of EMT-associated genes than CD24-negative cells [131]. The field is currently lacking well-designed studies that experimentally induce an EMT and thoroughly assess tumorigenicity and other CSC traits; however, there is no evidence to suggest that ovarian cancer is different than the other epithelial cancers where the EMT has been implicated in promoting CSC characteristics.

Immunosuppression Immune cell evasion is a hallmark characteristic of cancer: while various immune cells infiltrate the tumor microenvironment; they fail to launch effective attacks against the cancer [132]. The presence of immune cells within ovarian tumors is well-documented; however, the antitumor response is often negated by strong immunosuppressive signals [133,134]. These signals are also often associated with a worse prognosis for the patient. For example, patients with high intratumoral levels of FOXP3, a marker of immunosuppressive regulatory T-cells, have a dramatically reduced survival compared to those with low levels [135]. This immunosuppressive program is the net result of complex interactions between the immune system, the tumor cells themselves, and adjacent noncancer tissue. There is a strong relationship between immune cell regulation and the EMT. In fact, many of the EMT-

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inducing cytokines also serve as immunoregulators, including members of the TGFB family, various interleukins, and more. While there is a fair amount of work documenting the cooccurrence of the mesenchymal state and immunosuppressive environments in various cancers, more work is required to improve our understanding of how these phenomena arise and how they are maintained [136]. It is currently unclear to what extent the EMT promotes immunosuppression, how much the EMT is driven by cytokines secreted from immunosuppressive immune cells, or how additional factors such as stromal cells and vasculature contribute to both. It is likely the net effect of signals from many sources, including both the immune and cancer cells. The relationship between EMT and immunosuppression has yet to be studied in ovarian cancer; however, findings from other carcinomas may provide insight into its relevance. Various cytokines released from tumorinfiltrating leukocytes have been shown to promote EMT and metastasis [137], suggesting that the immune component contributes to carcinoma cell plasticity. Interestingly, the carcinoma cells themselves can also promote immunosuppression. When cocultured with splenocytes, both murine and human melanoma cells overexpressing the EMT-inducing transcription factor Snail cause, compared to control melanoma cells, reduced leukocyte proliferation, a lower proportion of CD4 + and CD8 + T-cells, and enhanced differentiation of immunosuppressive regulatory T cells and dendritic cells [138]. Additionally, inhibiting Snail with an intratumoral injection of small-interfering RNAs abrogated immunosuppression, leading to an enhanced infiltration of CD4+ and CD8+ T-cells and impaired tumor growth [138]. The mechanisms promoting the immunosuppressive phenotype are less clear; however, most findings point to the secretion of immunosuppressive cytokines. In a lung adenocarcinoma model, tumor cell secretion of TGFB has been shown to skew tumor-associated macrophages toward an anti-inflammatory “M2” phenotype by inhibiting TNF signaling through the interleukin receptor-associated kinase (IRAK)-M [139]. Another study has shown that miR-200 and Zeb1 regulate programmed death-ligand 1 (PD-L1) expression in lung cancer cells, with the mesenchymal state being associated with elevated PD-L1 secretion, causing suppression of CD8+ T-cells, which could be alleviated by treatment with an anti-PD-L1 antibody [140]. A recent study using breast carcinoma xenograft models has demonstrated that injecting flow cytometry-enriched EPCAMlo (mesenchymal) cells lead to immunosuppressive tumors relative to EPCAMhi (epithelial) cells [141]. Also, injecting various mixtures of these epithelial and mesenchymal cells results in comparable immunosuppressive phenotypes, suggesting that even a relatively small population of mesenchymal cells [ 10%] can program the majority of the tumor

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FIG. 3 A schematic representation of the consequences of the EMT on ovarian cancer progression. The transition has been associated with cancer metastasis, chemoresistance, and immunosuppression through complex interactions with the immune system. EMT, epithelial-to-mesenchymal transition; MET, mesenchymal-to-epithelial transition.

Carboplatin

Apoptosis

Chemoresistance MET

EMT

Metastasis / invasion Regulatory dendritic cells Regulatory T-cells

Immunosuppression

CD8+ T-cells

M2 macrophages

microenvironment, protecting the epithelial component from the antitumor response [141] (Fig. 3).

A THERAPEUTIC OUTLOOK: TARGETING PLASTICITY Given the collection of procancer traits regulated by the EMT, therapeutic control over carcinoma cell plasticity may yield tremendous benefits. Indeed, many have discussed developing therapies tailored to the molecular subtype of a given cancer [20,70,71]; however, discussion has focused on choosing therapies based on sensitivities of subtypes, rather than attempting to reprogram the cancer itself. Targeting the sensitivities of subtypes would likely improve outcomes over conventional therapy; however, given the heterogeneity within tumors, it seems probable that not all cells would be equally sensitive, leading to tumor recurrence. A therapy based on reprogramming the more-mesenchymal carcinoma cells to an epithelial state could render the tumor less metastatic and more susceptible to platinum-based chemotherapy. It may also activate the immune system, promoting an antitumor response, as was observed in the abovementioned study that found an intratumoral injection of small-interfering RNAs against Snail lead to immune activation within the tumor [138]. Ultimately, therapeutic reprogramming may be an effective complement to other treatments.

Occasionally, it is possible to collect repeated samples from a tumor—before and after a chemotherapy regimen, for example—but plasticity is always confounded with compositional differences, such as the enrichment of a population of chemoresistant cells, that were present, but proportionally rare before treatment. It is clear that ovarian cancer cells display plasticity when experimentally forced to do so, but then again, skin cells can be reprogrammed into pluripotent stem cells with the right experiment. So the question remains, do ovarian carcinomas display plasticity in their natural setting? Given the number of signaling pathways that seem to regulate the EMT, it seems reasonable to assume that the cancer cells do respond to changes in the tumor microenvironment, which likely alter the milieu of signaling ligands. The above-mentioned study [110] that described an induction of an EMT gene expression signature when ovarian cancer cells were transferred from adherent to suspension cultures is promising because it did not involve ectopically expressing EMT genes and demonstrated that these cells can dynamically respond to their environment. Moving forward, genetic lineage tracing experiments to permanently label cells upon the induction of an EMT, followed by functional selection such as metastasis or chemotherapy treatment of tumor models will greatly improve our understanding of plasticity in the context of tumor progression. Until then, while we know ovarian carcinoma cells are capable of remarkable plasticity, the assumption that plasticity plays a role in tumor progression is strictly hypothetical.

Does Plasticity Actually Play a Role in Tumor Progression? Plasticity, by its strictest definition, is difficult to observe in the context of cancer, as it requires the tracking of an individual cell’s phenotype over time. Often, we can only interrogate tumors by collecting molecular and visual data as snapshots of the moment the sample was collected. Temporal information is rarely acquired.

CONCLUSION Cell plasticity is critical for many biological processes, from embryo development and tissue homeostasis to cancer. Following each ovulation, the OSE relies on its plasticity to undergo an EMT and repair the ovulatory wound. Likewise, the FTE relies on this plasticity to

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REFERENCES

maintain homeostasis. This same plasticity is thought to contribute to some of the most detrimental features of ovarian cancer, including metastasis and the resistance to chemotherapy. Improving our understanding of the molecular determinants of this phenomenon would provide valuable insight into ovarian homeostasis and could potentially improve personalized medicine: new subtypes of ovarian cancer could be defined, targeted therapies for each subtype could be developed, and perhaps plasticity itself could be targeted, steering a cancer to a more-manageable state. Discussion of epithelial cell plasticity often revolves around the notion of a cell that is capable of transition between epithelial and mesenchymal states, as it has been described in this chapter. This focus is likely the consequence of the transition being readily observable and resulting in remarkable functional changes. However, it should be recognized that cellular plasticity is not limited to this transition and likely comprises responses to many transient stimuli, such as inflammation, hypoxia, circadian rhythm, and more. Environmental changes throughout the reproductive cycle beyond ovulatory wound repair may also drive transient responses in the OSE and FTE that may be of interest for understanding ovarian homeostasis and tumorigenesis. As technology improves and methodologies are developed to assess broadly cell state in vivo, it will become increasingly possible to determine the effect of these natural stimuli on the OSE, FTE, and other tissues of the ovary. Only then will we be able to grasp fully the complexities of this dynamic organ.

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C H A P T E R

34 Novel Therapeutic Approaches and Targets for Ovarian Cancer Rebecca A. Previs, Gordon B. Mills, Shannon N. Westin Abbreviations APC CDK CTLA-4 EGF EGFR ERK EZH2 FGF HDAC HIF HRG JNK KIT LOH MAPK MEK MHC NRG PARP PD-1 PDGF PDL-1 PlGF PI3K PRC2 RAF RAS RECIST Rheb-GTP SCCOHT SWI/SNF TCGA TGF-α TSC VEGF

treatment of epithelial ovarian cancer [1,2]. Although the majority of epithelial ovarian cancers respond to chemotherapy initially; unfortunately, the majority of these tumors will recur. With the emergence of drug resistance to primary- and secondary-line agents, the chance to offer a cure diminishes [3]. Traditional approaches to treatment have relied on a “one size fits all” approach with cytotoxic therapies being the mainstay for all histologic subtypes of epithelial ovarian cancer. A better understanding of cancer biology, coupled with advances in sequencing, and the availability of targeted and immunotherapy provide opportunities for precision medicine, which is tailoring a specific treatment for a particular cancer type at a precise time point.

antigen-presenting cell cyclin-dependent kinase cytotoxic T lymphocyte antigen-4 epidermal growth factor epidermal growth factor receptor extracellular signal-regulated kinase enhancer of zeste homolog 2 fibroblast growth factor histone deacetylase hypoxia-induced factor heregulin c-Jun N-terminal kinase stem cell factor loss of heterozygosity mitogen-activated protein MAPK-ERK kinase major histocompatibility complex neuregulin poly-ADP-ribose polymerase programmed cell death 1 platelet-derived growth factor programmed cell death ligand 1 placental growth factor phosphatidylinositol 3-kinase polycomb repressive complex 2 v-raf 1 murine leukemia viral oncogene homolog 1 retrovirus-associated DNA sequences response evaluation criteria in solid tumors Ras homolog enriched in brain-GTP small cell carcinoma of the ovary, hypercalcemic type SWItch/sucrose nonfermentable The Cancer Genome Atlas transforming growth factor-α tuberous sclerosis complex vascular endothelial growth factor

MOLECULAR ABERRATIONS IN OVARIAN CANCER

INTRODUCTION Surgical management with primary cytoreduction and adjuvant chemotherapy with carboplatin and paclitaxel remain the mainstays for upfront management and The Ovary https://doi.org/10.1016/B978-0-12-813209-8.00034-0

Ovarian cancer is a diverse disease, clinically, pathologically, and molecularly. While the majority of tumors are epithelial in nature, including serous, endometrioid, mucinous, clear cell, Brenner, undifferentiated, and carcinosarcomas, other more rare histologies include sex cord stromal tumors (comprised of Sertoli-Leydig, granulosatheca cell tumors, and lipid cell tumors) and ovarian germ cell tumors (choriocarcinomas, dygerminomas, immature teratomas, yolk sac, and embryonal). While epithelial ovarian carcinomas have been traditionally treated with the same therapeutic algorithm in the past, growing evidence suggests that the unique molecular profile that differentiates these tumors should be considered [4]. Initiatives such as The Cancer Genome Atlas (TCGA) and the International Cancer Genomics Consortium have completed large-scale studies of multiple tumor types, including ovarian cancer, to characterize alterations within pathways. TCGA evaluated 489 high-grade serous ovarian cancers and identified four subtypes: immunoreactive,

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differentiated, proliferative, and mesenchymal [5]. Findings from this analysis provided insight into the types of mutations, copy number alterations, and gene expression within this histologic subtype. Almost all were purely high-grade serous samples and contained a TP53 mutation and frequent germline or somatic mutations in BRCA1, BRCA2, and/or other homologous recombination (HR) DNA repair genes. However, other than these aberrations, there were few recurrent mutations in other genes or pathways with high-grade serous ovarian cancer being primarily driven by DNA copy number aberrations. While the majority of alterations identified are not currently actionable, large studies like this one inform future directions for development of targeted agents. Molecular aberrations that characterize the other histologies are listed in Table 1. Low-grade serous ovarian cancers are characterized by mutations in the retrovirus-associated DNA sequences v-raf 1 murine leukemia viral oncogene homolog 1 (RAS/RAF) pathway and in the phosphatidylinositol 3-kinase (PI3K/AKT). Mucinous tumors frequently possess KRAS mutations, which have been classified as “driver” mutations [6,7] and are potentially actionable. However, targeting the RAS/RAF pathway in ovarian cancer with monotherapy has only demonstrated modest responses. Like lowgrade serous ovarian cancer, clear cell carcinomas harbor a number of mutations within the PI3K/AKT pathway [8–10] but also contain ARID1A mutations and have overexpression of the interleukin 6-signal transducer, which serves as an activator of transcription 3-hypoxia-induced factor (IL6-STAT3-HIF) pathway [11,12]. Similarly, endometrioid ovarian cancers contain aberrations within both the PI3K/AKT and RAS/RAF pathways [13,14]. Other mutations that have been described but are not yet actionable include FOXL2 [15] and DICER1 [16,17] mutations in granulosa cell tumors and Sertoli-Leydig tumors, respectively. Small cell carcinoma of the ovary, hypercalcemic type (SCCOHT), has been characterized by a deleterious germline or somatic inactivating mutations in the SWItch/sucrose nonfermentable (SWI/SNF) chromatin-remodeling complex in SMARCA4, also known as BRG1 [18–20]. Loss of SMARCA4 and SMARCA2 protein expression has been found to be both sensitive and specific for this rare type of ovarian cancer [19,21]. Interestingly, ARID1A, which is mutated in small cell ovarian cancers and SMARCA4, play highly related roles in the SWI/SNF complex and in DNA repair. Although, these mutations cannot currently be targeted directly, SMARCA4 and ARID1A mutations may signal sensitivity to poly-ADP-ribose polymerase pathway (PARP) inhibitors, and these mutations are being evaluated as potential biomarkers in clinical trials. Historically, histopathology has affirmed the marked phenotypic heterogeneity within ovarian cancers. As with all therapies directed at tumors, intratumoral

TABLE

Epithelial

1 Molecular Characteristics Neoplasms

Cancer

Histology

Characteristics

Percentage (%)

Low-grade serous

TP53 mutation

7
10

KRAS mutation

19
35

BRAF mutation

2

EGFR amplification

17
73

PIK3CA amplification

20

PIK3CA mutation

2

PTEN mutation

17
33

PIK3CA mutation

20

BRAF mutation

4

TP53 mutation

10
50

HER2 amplification

14
41

PTEN mutation

3
5

PIK3CA mutation

20
34

PIK3CA amplification

20

AKT amplification

14

EGFR amplification

50

EGFR mutation

15

KRAS mutation

40
60

BRAF mutation

10

HER2 amplification

45

EGFR mutation

66

TP53 mutation

80
96

BRCA1/2

10
20

KRAS mutation

0
12

PIK3CA amplification

68

Clear cell

Mucinous

High-grade serous

Other

Ovarian

Classification

Endometrioid

Sex cordstromal

of

Molecular

Adult granulosa FOXL2 mutation

50
89

Sertoli-Leydig

DICER1 mutation

0
100

Small cell carcinoma of ovary, hypercalcemic type

SMARCA4 mutation

94

heterogeneity and the tumor microenvironment must also be recognized as contributing to efficacy. The mutational profile of the primary tumor may not match the profile of a metastasis. Metastasized tumor cells evolve and acquire additional mutations, some of which may contribute to resistance to therapies [22,23]. As a better understanding of cancer cell biology emerges as

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described by Hanahan and Weinberg in their seminal papers, the 10 hallmarks of cancer must be considered in tumorigenesis and how newer therapies affect the growth and metastasis of cancer cells [24]. While traditional cytotoxic therapies primarily target the division of cancer cells, chemotherapy also kills rapidly proliferating normal cells within the gastrointestinal, hematopoietic, and other body systems. Ideally, targeted therapy would match identified alterations within the tumor itself thus conferring high efficacy and limited toxicity. Alternatively, targeted agents could disrupt the growth of the cancer within the microenvironment and prevent further metastasis or block the growth of existing metastasis. This would spare normal tissues and avoid the untoward side effects patients experience during their cancer treatment. In the treatment of ovarian cancer, PARP inhibitors represent one of the greatest successes in biomarker directed therapy with the recent approval of three different PARP inhibitors, which show marked activity in tumors with abnormalities in the homlogous recombination (HR) pathway mediated by the BRCA1/2 tumor suppressor genes. Other targets for ovarian cancer patients include the immune system, TP53, angiogenesis, altered molecular pathways, and aberrant cell-surface molecules. These novel therapeutic approaches offer patients hope for additional treatment options in the recurrent setting. While the field of targeted therapy evolves rapidly, the objective of this chapter is to provide a broad overview of relevant pathways in ovarian cancer and the targeted therapies that are currently being studied in clinical trials.

HOMOLOGOUS RECOMBINATION AND POLY-ADP-RIBOSE POLYMERASE INHIBITORS Homologous recombination (HR) is the dominant and importantly most efficient DNA repair pathway for double-strand DNA breaks. In patients harboring a BRCA1/2 mutation, the base excision repair, single DNA strand break pathway and the double-stranded DNA repair nonhomologous end-joining pathways play more important roles in DNA repair. This dependency upon double-strand break repair mechanisms leads to the enhanced activity of poly-ADP-ribose polymerase (PARP) inhibitors that block base excision repair and to a degree, nonhomologous end-joining, in cells that have deficient or mutated BRCA1/2 [25,26] (Fig. 1). Once activated, PARP binds to DNA and induces a structural change, which is the rate-limiting step in the base excision repair pathway. This conformational change then recruits repair proteins to the site of damage. PARP inhibitors were introduced to the clinic in

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2005, following preclinical work that suggested that BRCA1- and BRCA2-deficient cells were up to 1000fold more sensitive to PARP inhibition than wild-type cells in vitro [25]. The HR pathway is complex, but the mechanism of PARP inhibitors is multifold and induces synthetic lethality with defects in HR [25,27]. The side effect profile varies slightly depending on the PARP inhibitor used and whether it is in combination with other therapeutics. Common side effects of PARP inhibition including nausea, gastrointestinal upset, fatigue, anemia, and myelosuppression. If PARP inhibitors are used in combination with chemotherapy, more myelosuppression may be encountered, requiring close monitoring and dose reductions. Over a decade later, three PARP inhibitors have been approved by the Food and Drug Administration (FDA) for use in ovarian cancer patients with recurrent disease (Table 2). The effectiveness of PARP inhibitors in BRCAmutated tumors is due to several other important mechanisms. PARP triggers apoptosis, in part, through repair of DNA double-strand breaks and the low fidelity nonhomologous end-joining repair pathways. This occurs due to the accumulation of errors in the DNA that ultimately cannot be repaired [28]. Another mechanism of action includes PARP trapping, which is the ability of a PARP inhibitor to trap PARP1 and PARP2 at the site of damage within the DNA. The complex that forms is cytotoxic and interferes with DNA replication [29]. PARP inhibitors vary in their DNA trapping activity, with veliparib having the lowest trapping activity and talazoparib the highest activity. The relative roles of inhibition of PARP enzyme activity and of DNA trapping in the efficacy of PARP inhibitors in ovarian cancer remain to be fully elucidated. Multiple potential mechanisms of resistance to PARP inhibitors have been identified, primarily, in model systems. These fall into three major categories: (1) those that reconstitute HR competence including healing of mutations or reexpression of HR pathway members such as BRCA1/2 or downregulation of 53BP1, PTIP, or PARG; (2) those that cause replication fork protection such as decreased proliferation or loss of enhancer of zeste homolog 2 (EZH2) or MLL3/4; and (3) those that alter function of PARP such as increased export of PARP through ABC transporters, PARP loss or mutation or loss of SLFN1 or CHD1. Examples are presented in Fig. 1B. As indicated below, most PARP inhibitor studies in ovarian cancer have focused on monotherapy in the maintenance setting. However, a number of combination trials such as PARP inhibitors combined with smallmolecule inhibitors of DNA damage checkpoints, MEK, CDK4/6, and angiogenesis as well as with immune oncology agents are ongoing. The combination of PARP inhibitors with antiangiogenic agents such as cedirinib is very promising in early trial data.

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34. NOVEL THERAPIES IN OVARIAN CANCER

BRCA-mutated cell No homologous recombination

Single-strand break PARP

Cell death

PARP inhibitors

Cell survival and DNA repair

Normal cell

(A)

BRCA1 BRCA2

↓53BP1

A secondary BRCA mutation restores BRCA function

↓53BP1 or absent 53BP1 partially restores homologous recombination repair

Overexpression of P-glycoprotein increases effux of PARP inhibitors

(B) FIG. 1 (A) The role of PARP and DNA damage repair. PARP binds to single-strand breaks in DNA and in normal cells, other enzymes are recruited, and the damage is repaired. PARP inhibitors induce double-strand DNA breaks, which is repaired in non-BRCA-mutated cells by homologous recombination. In cells that have a mutation in BRCA1 or BRCA2, two genes that encode components of the homologous recombination pathway, the DNA damage cannot be sufficiently repaired, and synthetic lethality is induced. Cell cycle arrest occurs, and the cell undergoes apoptosis. (B) Mechanisms of resistance to PARP inhibitors. Three mechanisms of acquired resistance to PARP inhibitors in BRCA-mutated tumors include secondary molecular defects that restore BRCA function, reduced or absent 53BP1, which is a chromatin-associated factor that promotes immunoglobulin class switching and DNA double-strand break repair by nonhomologous end joining, and overexpression of P-glycoprotein, which increases the efflux of PARP inhibitors out of the cell. PARP, poly-ADP-ribose polymerase.

TABLE 2

Clinical Trials Treating Evaluating Monotherapy PARP Inhibitors in the Treatment of Ovarian Cancer Patients Indication

BRCA Status

N

Response Rate (%)

Duration of Response (months)

3 lines chemotherapy

Germline mutation

137

34

7.9

All lines

Germline mutation

300

36

7.4

Rucaparib (Rubraca)

2 lines of chemotherapy

Germline/somatic mutation

106

54

9.2

Niraparib (Zejula)

All lines

Germline mutation

20

40

12.9

All lines

No mutation

22

9

NR

Up to 3 prior lines

Germline mutation

50

26

NR

Olaparib (Lynparza)

Veliparib (ABT-888)

BID, twice daily; NR, not reported; PO, by mouth.

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Olaparib BRCA1/2 mutations are the only approved molecular targets for ovarian cancer patients with an associated biomarker. Olaparib is the most studied PARP inhibitor. A phase 2 study was initiated following activity of olaparib demonstrated in a phase I study [30]. This multicenter trial enrolled patients with BRCA1/2 mutations that had been treated with at least one previous line of therapy to receive either continuous olaparib dosed at 100 mg twice daily or 400 mg twice daily. The primary endpoint for this study was objective response rate, and the group that received the lower dose achieved a response rate of 12.5%. The group that received the 400 mg twice daily dose had higher levels of response with an objective response rate of 33% and a clinical benefit rate of 57.6% [31]. Multiple phase 2 studies in recurrent ovarian highgrade serous carcinoma have been performed and confirmed favorable responses to olaparib in patients with and without BRCA mutations [31–36]. Olaparib was the first PARP inhibitor to be approved by the FDA on December 19, 2014, for use in recurrent ovarian cancer patients with a BRCA1/2 mutation, who had received three or more prior regimens based on pooled data from early studies. In a phase 2, randomized study, 162 patients with recurrent platinum-sensitive ovarian cancer were enrolled to receive olaparib plus carboplatin and paclitaxel followed by olaparib monotherapy as maintenance or carboplatin and paclitaxel alone [32]. While there was no significant difference in response rates, the addition of olaparib to standard chemotherapy significantly improved progression-free survival (PFS), with a median of 12.2 months (compared to 9.6 months in the standard therapy arm). Importantly, this observation was more pronounced in patients with a known BRCA mutation and became more pronounced during the maintenance phase of the study. Use of olaparib as a maintenance therapy has also been evaluated in the platinum-sensitive setting. In a phase 2 trial, patients with platinum-sensitive, recurrent disease were treated with olaparib or placebo until disease progression after they achieved a complete or partial response to previous platinum-based therapy. This study did not exclude patients on the basis of their BRCA mutation status and noted a significant improvement in PFS. In a subgroup analysis, treatment of patients with a BRCA 1/2 mutation with olaparib had a dramatic effect on PFS, despite not reaching significance for overall survival (OS) in the initial analysis. Of note, the European Medicines Agency and FDA has approved olaparib for use as a monotherapy as a maintenance treatment of platinum-sensitive patients with a BRCA mutation and recurrent disease, who have previously responded to platinum-based chemotherapy [34,37]. These data were

551

recently confirmed in the SOLO-2 study, which randomized patients with germline BRCA mutation to olaparib vs placebo. A significant improvement in PFS was observed in those patients treated with olaparib [38]. In platinum-resistant populations, objective response rates have ranged from 33% to 42% [35,39–41].

Rucaparib Rucaparib (CO-338) is also a potent small molecular inhibitor of both PARP1 and PARP2 at nanomolar concentrations [42]. Phase 1 and 2 studies of intravenous (IV) rucaparib were evaluated in combination with temzolomide and no serious side adverse events were reported [43,44]. A phase 2, open label multicenter trial included BRCA1 and BRCA2 mutation carriers with advanced breast and/or ovarian cancer. Both IV and oral administrations of rucaparib were assessed to determine the safety, tolerability, dose-limiting toxicities, pharmacodynamics, and pharmacokinetic profiles. The patients who received IV rucaparib demonstrated an overall objective response rate of only 2%, but 41% of patients achieved stable disease for 12 weeks, with three patients having stable disease for greater than a year. The overall response rate for oral rucaparib, across all of the dosing levels, was 15%. The conclusion from this study was that the drug is well tolerated and resulted in target inhibition in target tissues even at the lowest dosing schedules. Continuous dosing was required for optimal response [45]. ARIEL2 is a multicenter, two part, phase 2 trial that included patients with recurrent, platinum-sensitive high-grade ovarian carcinoma. In this study, the patients were classified into three groups based on blood- and tissue-based assay: BRCA mutant tumors (germline or somatic); BRCA wild type and homologous recombination defective (HRD) based on loss of heterozygosity (LOH) high; or BRCA wild type and LOH low. Patients were treated with 600 mg twice daily until progression or discontinuation. This study demonstrated that patients who are treated with rucaparib and have a BRCA mutation or have HRD defined by LOH and are BRCA wild type have longer PFS than patients who are BRCA wild type and have LOH low carcinomas. Further data are being collected to better identify patients with HRD. These findings suggested the efficacy of PARP inhibitors beyond patients with BRCA mutations. On December 19, 2016, the FDA granted accelerated approval to rucaparib in patients with BRCA mutated (germline or somatic) advanced ovarian cancer who have been treated with two or more lines of chemotherapy. Other ongoing phase 3 trials include ARIEL3, which is study of rucaparib as switch maintenance following platinum-based chemotherapy in platinum-sensitive, high-grade serous, or endometrioid ovarian cancer

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34. NOVEL THERAPIES IN OVARIAN CANCER

similar to the SOLO-2 and NOVA (see below) studies. Primary outcome measures are disease progression according to response evaluation criteria in solid tumors (RECIST). Patients enrolled into this study are stratified into three groups based on mutation status to evaluate patients’ response to maintenance treatment with rucaparib vs placebo (NCT01968213). Following this trial, the FDA approved rucaparib in 2018 as maintenance therapy for women who are in complete or partial response to platinum-based therapy. ARIEL4 is a phase 3 trial-evaluating patients with recurrent, BRCA mutant, high-grade ovarian cancer. In this study, patients either receive rucaparib or chemotherapy (monotherapy platinum or platinum-based double chemotherapy). The primary outcome is PFS.

Niraparib Early success in phase 1 trials was observed in patients with advanced solid malignancies in patients treated with niraparib. In part A of the Niraparib phase I study, cohorts were enriched for BRCA1 and BRCA2 mutation carriers and received escalating doses. Eight of 20 patients with a BRCA1 or BRCA2 mutation and ovarian cancer had partial responses. Two of four BRCA mutation carriers with breast cancer had partial responses. Antitumor activity was also reported in sporadic high-grade serous ovarian cancer, non-small-cell lung cancer, and prostate cancer. A recommended phase 2 dose of 300 mg/day was recommended [46]. Recently, the results of the phase 3 NOVA trial of niraparib successfully achieved its primary endpoint of PFS regardless of BRCA mutation or HRD status [47]. This was a doubleblind placebo-controlled trial that included over 500 patients with recurrent ovarian cancer that were platinum-sensitive. In the patients with a germline BRCA mutation who received niraparib, there was a 21-month median PFS compared to the 5.5 months in the placebo group. In patients who did not have a germline mutation, there was also a PFS benefit for patients received niraparib over placebo (12.9 vs 3.8 months) [47]. This effect was attenuated in patients without BRCA mutation or evidence of HRD (6 vs 3 months). These findings led to the US FDA’s approval of niraparib for the maintenance treatment of all patients with recurrent epithelial ovarian, fallopian tube, or primary peritoneal carcinoma who are in complete or partial response to their most recent platinum-based chemotherapy. However, whether the magnitude and duration of benefit in the BRCA wild type and HRD normal tumors warrants therapy with niraparib remains controversial. Niraparib is under evaluation for primary maintenance and in patients with multiple lines of therapy as well. QUADRA is an ongoing phase 2, open-label,

single-arm study evaluating the efficacy and safety of niraparib in ovarian cancer patients who have received three or four previous chemotherapy regimens with a primary outcome measure of the activity of niraparib (NCT02354586). PRIMA is a phase 3, randomized, double-blind, placebo-controlled trial in patients with ovarian cancer who have homologous recombinantdeficient tumors. Patients are randomized 2:1 (niraparib: placebo) after at least four cycles of a platinum-based chemotherapy, must have achieved a complete or partial response (with no tumor greater than 2 cm) to a platinum-based regimen, and must have either a normal CA125 or a CA125 that has decreased to more than 90% during their frontline therapy. Primary outcome measure is PFS, and secondary outcome measures include OS, safety and tolerability of niraparib, time to progression on the next anticancer therapy, and patient reported outcomes (NCT02655016).

Veliparib Veliparib (ABT-888) is a small molecule that inhibits PARP1 and PARP2 at nanomolar concentrations [48]. It has good bioavailability and has been shown to cross the blood-brain barrier. In preclinical syngeneic and xenograft models, veliparib has been shown to potentiate the effect of platinum-based therapy, cyclophosphamide, radiation, and temzolomide [48]. Most early clinical studies have evaluated veliparib in combination with cytotoxic chemotherapy including the I-SPY2 breast cancer trial, which evaluated the combination of veliparib and carboplatin in triple negative breast cancer and had several patients with objective responses [34]. The ability to combine veliparib with chemotherapy may be due in part to its low PARP trapping activity. Early phase 1 studies have demonstrated the maximum tolerated dose to be 400 mg twice daily [49–51]. To evaluate veliparib in a population of BRCA mutationcarrying women with recurrent ovarian cancer, an open label, phase 2, multicenter clinical trial was conducted [52]. In this study, up to three prior therapies were allowed; 60% of patients were platinum resistant. One grade 4 event was reported: thrombocytopenia. Grade 3 events that occurred were fatigue, nausea, leukopenia, neutropenia, dehydration, and elevations in ALT. The proportion of patients responding was 26% (90% CI [confidence interval]: 16%
38%), and the median PFS was 8.18months. For platinum-resistant and platinumsensitive patients, the proportion responding was 20% and 35%, respectively. Phase 1 data of oral cyclophosphamide in combination with veliparib showed tolerability of the combination and clinical activity in patients with BRCA mutant

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TARGETING ANGIOGENESIS: VASCULAR ENDOTHELIAL GROWTH FACTOR AND VEGF RECEPTORS

tumors, with 6 of 13 patients having a partial response and 3 additional patients with prolonged disease stabilization [49]. A phase 2 trial in pretreated patients with BRCA mutations evaluated the effect of veliparib in combination with oral cyclophosphamide in high-grade serous ovarian, fallopian tube and peritoneal cancers. Patients were randomized to receive either cyclophosphamide alone (50-mg orally daily) or in combination with veliparib (60-mg orally daily) in 21-day cycles with crossover to the combination group allowed. Seventytwo patients were evaluable, and one complete response was achieved in each arm with three and six partial responses achieved in the combination arm and cyclophosphamide alone arm, respectively. Although well tolerated, there was no improvement in the response rates or PFS with the addition of low dose veliparib in combination with cyclophosphamide [53]. Other reports of clinical benefit of veliparib have been reported or are ongoing. One phase 1/2 study demonstrated an overall response rate of 65.7%; the progression-free and OS in the intention-to-treat population was 5.5 months (4.9–7.3 months, 95% CI) and 15.2 months (10–17.3 months, 95% CI), respectively [54]. In addition to fatigue and gastrointestinal toxicities, PARP inhibitors are well known to cause hematologic toxicities such as anemia and thrombocytopenia which have limited success in chemotherapy combinations, but veliparib has been combined with standard chemotherapy without significant dose reductions of either agent [55]. GOG-3005 is an ongoing phase 3 placebo controlled study of carboplatin and paclitaxel with or without concurrent and continuation maintenance veliparib in patients with previously untreated Stages III or IV high-grade serous ovarian carcinoma (NCT02470585). This study includes three experimental arms. All patients receive carboplatin and paclitaxel for six 21-day cycles. Arm one includes placebo as a part of the frontline chemotherapy and is followed by placebo maintenance therapy for an additional 21-day cycles. In arm two, patients receive veliparib plus chemotherapy for six cycles followed by placebo maintenance. In arm three, patients receive veliparib plus chemotherapy for six cycles followed by veliparib maintenance. Primary outcome measures include PFS, and secondary outcome measures are OS and disease-related symptom scores.

Talazoparib Talazoparib (BMN673) is the most active of the PARP inhibitors in clinical evaluation both in terms of its inhibition of PARP enzyme activity and PARP trapping activity. This inhibitor has shown intriguing activity in a number of diseases. Its development program in ovarian cancer is likely to focus on combination therapy with targeted and immuno-oncology agents.

553

TARGETING ANGIOGENESIS: VASCULAR ENDOTHELIAL GROWTH FACTOR AND VEGF RECEPTORS Angiogenesis is a complex process regulated by numerous pro- and antiangiogenic factors [56,57]. Normal ovarian physiology relies on angiogenesis for the transport of nutrients, growth factors, and oxygen particularly following ovulation. However, ovarian cancer growth, progression, and metastatic spread rely on the process of neovascularization [58]. This angiogenic switch, which has been described in solid malignancies, leads to the formation and growth of new vasculature. This vasculature tends to be abnormal and have thinner walled basement membranes, which allows for increased permeability [59,60]. Tumor cells secrete proangiogenic factors such as vascular endothelial growth factor (VEGF), platelet-derived growth factor (PDGF), placental growth factor (PlGF), and fibroblast growth factor (FRF), which stimulate endothelial cells to proliferate and migrate. Stromal cells within the tumor microenvironment can also release factors that further contribute to the angiogenic switch. The VEGF family of growth factors includes VEGF-A, VEGF-B, VEGF-C, VEGF-D, VEGF-E, which bind to VEGF receptors: VEGFR1 (Flt1), VEGFR2 (Flk-1), and VEGFR3 (Fig. 2). Expression of VEGF and VEGFRs has been found to be higher in ovarian cancer tissue than normal tissue making this process and pathway an exciting target for ovarian cancer. Bevacizumab is a VEGF humanized monoclonal antibody that has been widely studied in ovarian cancer in combination with carboplatin and paclitaxel, as a maintenance therapy, and in the recurrent setting in patients with platinum-sensitive and platinum-resistant disease. ICON7 was a phase 3 trial that assigned ovarian cancer patients to receive carboplatin and paclitaxel every 3 weeks for six cycles with or without bevacizumab. In patients who were randomized to receive bevacizumab, they received it concurrently and then as a maintenance treatment for 12 additional cycles (or until disease progression). In an updated analysis of this study, PFS at 42 months was 22.4 months with carboplatin and paclitaxel alone and 24.1 months with the addition of bevacizumab (P ¼ 0.04). Patients who had higher risk disease (as defined by suboptimal cytoreduction or Stage IV disease) had a greater benefit with bevacizumab (PFS was 14.5 months vs 18.1 months and median OS was 28.8 vs 36.6 months, favoring the bevacizumab-treated arm) [61]. GOG 218 also evaluated bevacizumab added to standard frontline therapy in a phase 3 trial of advanced stage ovarian cancer patients. Patients received one of three treatments, all of which included six cycles of carboplatin and paclitaxel. Arm 1 received placebo combined with chemotherapy followed by placebo maintenance; arm 2 received bevacizumab with chemotherapy

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34. NOVEL THERAPIES IN OVARIAN CANCER

FIG. 2 Overview of ligands, receptors, interactions, and inhibitors of the angiogenesis pathway. Endothelial cells express the VEGF tyrosine kinase receptors (VEGFR1, VEGFR2, and VEGFR3). Tumor-derived VEGF-A interacts with VEGFR1 and VEGFR2. PlGF and VEGF-B bind to VEGFR1. VEGF-C, VEGF-D, and VEGF-E interact with VEGFR2 and VEGFR3. Also shown are PDGF receptor (PDGFR) and FGF rectors (FRFR) with their corresponding ligands, PDGF and FGF, respectively. The Tie2 tyrosine kinase binds ligands Ang1 and Ang2. Current agents being developed to disrupt these interactions are listed.

during cycles two through six followed by placebo maintenance; and arm 3 received bevacizumab with chemotherapy during cycles two through six followed by bevacizumab maintenance. Median PFS was 10.3, 11.2, and 14.1 months, respectively, with the most significant improvement in the arm including bevacizumab maintenance [62]. Bevacizumab has recently been approved by the FDA in the frontline setting. In the recurrent setting, patients with platinumsensitive recurrent ovarian cancer were randomized to receive carboplatin and gemcitabine with placebo or bevacizumab for 6–10 cycles followed by bevacizumab or placebo maintenance continued until disease progression. PFS in the bevacizumab arm was higher (12.4 vs 8.4 months) and there was a higher objective response rate and duration of response in the patients who received bevacizumab, however, there was no difference in OS (32.9 vs 33.6 months, P ¼ 0.65) although the study was not powered to detect a difference [63,64]. Similar results were observed in GOG 213 with a similar patient population. Patients were randomly assigned carboplatin and paclitaxel with or without bevacizumab. Bevacizumab was continued as maintenance until disease progression or toxicity. In addition to a statistically significant difference in PFS of 14 vs 10 months favoring the bevacizumab-treated arm (HR 0.61, 95% CI: 0.52
0.72), there was a trend in the same arm toward improved OS of 43 vs 37 months (HR 0.82, 95% CI: 0.68
0.996), which was significant upon reanalysis using corrected data [65]. This led to an FDA approval of

bevacizumab in combination with a platinum-based chemotherapy combination in platinum-sensitive recurrent ovarian cancer. Bevacizumab has also been approved in the platinumresistant setting in combination with paclitaxel, liposomal doxorubicin, or topotecan. This was based on the AURELIA phase 3 trial in recurrent, platinum-resistant ovarian cancer, which assigned patients to chemotherapy with weekly paclitaxel, pegylated liposomal doxorubicin, or topotecan with or without bevacizumab. Crossover to single-agent bevacizumab was permitted after progression if the patient received single agent chemotherapy alone. The median PFS was 3.4 months vs 6.7 months in the chemotherapy alone and chemotherapy with bevacizumab groups, respectively, but OS was not significantly improved [66]. The greatest different in PFS was seen in patients who received bevacizumab with weekly paclitaxel [67]. Bevacizumab has been evaluated in nonepithelial ovarian cancers such as recurrent granulosa cell tumors with a response rate of 38% and a clinical benefit rate of 63% [68]. Although bevacizumab has been evaluated in a variety of clinical settings, the patient population who best benefits and when they will derive the most benefit remains to be identified. Further, it is unclear if a patient who receives bevacizumab in the upfront setting will have as great a benefit in the recurrent setting. Importantly, bevacizumab has been associated with significant toxicity including bowel perforations and death. Thus in

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PHOSPHATIDYLINOSITOL-3-KINASE/AKT PATHWAY

the absence of biomarkers of benefit, the utility of bevacizumab and other angiogenesis inhibitors remains uncertain. However, as noted above, combinations with other targeted agents such as PARP inhibitors have demonstrated exciting preliminary results.

PHOSPHATIDYLINOSITOL-3-KINASE/ AKT PATHWAY The PI3K/AKT/mTOR pathway controls important normal cellular processes that are necessary for tumorigenesis and metastasis including cell survival, proliferation, regulation of cell cycle, angiogenesis, and metabolism [69]. Aberrations within this signaling pathway are one of the most common events in solid malignancies, especially ovarian cancer [70,71]. Alterations within this pathway are not limited to one histologic subtype, although specific abnormalities are more common in each subtype PI3K is a downstream effector of G-coupled protein receptors and tyrosine kinase receptors. PI3K is a heterodimer, consisting of a p110 catalytic and a p85 regulatory subunit. Activated PI3K phosphorylates phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2; PIP2) to generate phosphatidylinositol (3,4,5)-triphosphate (PI(3,4,5)P3; PIP3). AKT is then recruited to the cell membrane where it can be phosphorylated at a serine or threonine residue. The tumor suppressor, PTEN, and INPP4B are responsible for the negative regulation of activation. PTEN dephosphorylates PIP3, which produces PIP2 [69]. Constitutive activation of AKT can result from loss of PTEN or INPP4B or from AKT PH domain mutations. Activated AKT phosphorylates mTOR, which activates

p85 p110 IRS1

mTORC1. Indirectly, AKT can also activate mTORC1 via phosphorylation of the tuberous sclerosis complex (TSC2). Phosphorylated TSC2 then leads to the inactivation of the functional TSC1/TSC2 complex. When TSC1/ TSC2 is activated, TSC2 promotes the conversion of Ras homolog enriched in brain-GTP (Rheb-GTP) to RasGDP, which then inactivates mTORC1. Rheb-GTP is responsible for stimulating the activity of mTORC1 once TSC2 is phosphorylated or inactivated by AKT [72]. PI3K may be responsible for the activation of mTORC2, but the exact mechanism remains unclear [72] (Fig. 3). Activated mTORC2 phosphorylates a variety of other kinases, such as AKT and other kinases that regulate metabolism, lipogenesis, apoptosis, or the cytoskeleton. The drugs currently under investigation that target this pathway are primarily small molecular inhibitors of one or more pathway components. This pathway is involved in multiple feedback loops and cross talk with other pathways, necessitating the use of combination therapies for optimal efficacy. Aberrations within the PI3K/AKT/mTOR, such as PIK3CA amplification and/or PTEN loss, have led to promising preclinical work with mTORC1 inhibition [73]. Temsirolimus is an mTOR inhibitor that has been evaluated in multiple phase 1 and 2 studies of ovarian cancer patients. In patients with platinum-resistant, recurrent disease, 45 patients (22 ovarian cancer patients) were enrolled to receive temsirolimus treatment, however, despite being well-tolerated, the study did not meet predefined efficacy criteria. Several patients did have long-lasting PFSs (defined as greater than 7 months) [74]. In another phase 2 study of temsirolimus as a single agent, patients with measurable disease were enrolled to receive 25-mg IV

PIP3 AKT PIP2

PTEN

PDK1 mTORC2 TSC1/2

mTORC1

S6K

4E-BP1

FIG. 3

The PI3K/AKT signaling pathway. Activation of PI3K leads to downstream activation of AKT. Via phosphorylation events, AKT activates or inactivates other target molecules that are involved in the growth, survival, and proliferation of ovarian cancer cells. PI3K, phosphatidylinositol 3-kinase. VI. HUMAN OVARIAN CANCER

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temsirolimus weekly. Of 54 evaluable patients, 24.1% had a PFS that was greater than or equal to 6 months with only 9.3% experiencing a partial response [75]. Pilaralisib (GDC-094) is a selective inhibitor of the class I PI3Ks, and phase 1 studies have been completed showing that it was well tolerated. One patient with platinum-refractory epithelial ovarian cancer whose tumor had PTEN loss and PIK3CA amplification had a partial response to pilaralisib [76]. Preclinical evidence suggests that the PI3K/AKT pathway may impact resistance to carboplatin and paclitaxel [73]. The mechanism of this may occur from the phosphorylation of AKT, which promotes the survival of ovarian cancer cells via a reduction in P53 signaling or activation of downstream targets such as P70S6. Other PI3K inhibitors are being evaluated in combination with carboplatin and paclitaxel (NCT00756847). Preliminary evidence suggested that the combination was tolerable and clinical responses led to a dose expansion arm in ovarian cancer [77]. Preclinical work is currently ongoing looking at other agents including an mTORC1/2 inhibitor, AZD8055, which inhibited proliferation of ovarian clear cell and serous carcinomas in vitro [78]. Multiple AKT inhibitors have been studied in clinical trials. Perifosine, an AKT inhibitor, has been combined with docetaxel in patients with platinum and taxane resistant high-grade ovarian cancer, but median PFS and OS were 1.9 and 4.5 months, respectively, indicating limited activity [79]. One patient with a PTEN mutation achieved a partial response lasting 7.5 months. GSK2110183, an ATP-competitive inhibitor of the AKT 1, 2, and 3 isoforms, has been evaluated in multiple preclinical mouse models in combination with a MEK inhibitor. A dose finding study in platinum-resistant ovarian cancer in combination with carboplatin and paclitaxel is currently ongoing (NCT01653912). As with most small-molecule inhibitors, PI3K pathway inhibitors will likely demonstrate optimal activity only in combinations. Indeed, combinations of PARP inhibitors plus PI3K pathway inhibitors have demonstrated intriguing activity in phase 1 trials [80], and a number of phase 2 trials are ongoing with PI3K pathway and PARP inhibitors.

differentiation [81]. This pathway becomes activated after cell surface molecules activate RAS, which is a member of a family of GTPases, which then activates downstream RAF kinases. Downstream targets of the RAF kinases include mitogen-activated protein (MAPK)/ERK and MEK and ultimately, the activation of ERK leads to a variety of cell regulatory activities including metabolism, migration, and invasion [82,83] (Fig. 4). Cross talk between this pathway, the PI3K/AKT pathway, and mutually shared receptors, such as those within the epidermal growth factor receptor (EGFR) family, makes this pathway an attractive therapeutic target. Among epithelial ovarian cancers, low-grade serous histologies are characterized by MAPK mutations, which frequently leads to constitutive activation of this pathway [84,85]. Mucinous carcinoma, another type 1 ovarian carcinoma, has been reported to have activating KRAS mutations in up to 50% of cases [7]. Selumetinib, an inhibitor of MEK1/2, has been evaluated in a phase 2 trial of patients with low-grade serous ovarian cancer, and a 15% objective response rate was reported (one complete response and seven partial responses); 65% of patients had stable disease [86]. This has led to several clinical trials including the MILO study, which is comparing binimetinib (MEK162), another MEK inhibitor, with physician’s choice of chemotherapy (pegylated liposomal doxorubicin, paclitaxel, or topotecan) in patients with this disease (NCT01849874). Another study is currently investigating MEK162 in combination with weekly paclitaxel in patients with recurrent ovarian cancer (NCT01649336). Based on evidence that RAS mutations increase HR competency and MEK inhibitors induce HRD, we have recently instituted a clinical trial of olaparib and selumetinib (NCT03162627) in patients with low-grade serous tumors with KRAS mutations and in patients who have failed PARP inhibitors.

RAS

RAF

RETROVIRUS-ASSOCIATED DNA SEQUENCES/V-RAF 1 MURINE LEUKEMIA VIRAL ONCOGENE HOMOLOG 1 PATHWAY/MAPK-ERK KINASE/ EXTRACELLULAR SIGNAL-REGULATED KINASE The RAS/RAF pathway/MAPK-ERK kinase (MEK)/ extracellular signal-regulated kinase (ERK) pathway is a well-characterized signal transduction pathway that regulates cell growth, proliferation, survival, death, and

MEK1/2

ERK1/2

Binimetinib PD-0325901 Selumetinib Trametinib

FIG. 4 The RAS/RAF/MEK/ERK pathway and targeted therapies. Upon RAS activation, downstream RAF kinases are activated, leading to downstream cellular events involved in cellular metabolism, migration and invasion.

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EGFR PATHWAY TARGETS

EGFR PATHWAY TARGETS The EGFR signaling cascade is comprised of four receptors EGFR (ErbB-1), HER2/neu (ErbB-2), Her-3 (ErbB-3), and Her-4 (ErbB-4) that function by transmitting signals from the cell plasma membrane. Once activated by a specific ligand such as epidermal growth factor (EGF), transforming growth factor-α (TGF-α), or neuregulin (NRG), they undergo homodimerization (binding with the same receptor after ligand binding) or heterodimerization (binding with another receptor in the family). Once dimerization occurs, a series of phosphorylation events are initiated leading to downstream signaling in various cascades including the PI3K/AKT, MAPK/RAS, and c-Jun N-terminal kinase (JNK) pathways leading to transcription of genes involved in angiogenesis, cell proliferation, inhibition of apoptosis, and migration and invasion. EGFR is highly expressed in 35%
50% of ovarian cancers and has been implicated in cancer development [87,88]. Expression of EGF has been correlated with advanced stage in serous and clear cell ovarian cancers [89]. Overexpression of EGFR has been implicated in chemotherapy resistant cell lines in vitro making this pathway an attractive target to inhibit growth and metastasis. EGFR gene copy number has been associated with worse progression free and OS [4]. Gefitinib inhibits activation of EGFR through competitive binding of the ATP-binding domain. It has been explored in a variety of phase 2 trials ovarian cancer patients without promising clinical benefit. In a phase 2 GOG study, mutation in exons 18–21 of EGFR and/or immunohistochemical evidence of expression of EGFR were evaluated in patients enrolled on trial. In patients with EGFR-positive tumors, the response rate was 1 out of 11 patients (9%). EGFR expression was associated with a longer PFS [90]. Monotherapy gefitinib did not convey any clinical benefit despite successful pharmacodynamic endpoints seen after treatment, which included inhibition of the phosphorylation of EGFR in ovarian cancer tumor cells [91]. Combinations that have been reported or are ongoing include gefitinib with tamoxifen (no responses) [92], with anastrozole (NCT00181688), liposomal doxorubicin (active regimen but with significant skin toxicity) [93], topotecan (NCT00317772), and carboplatin and paclitaxel. In the phase 2 study that evaluated gefitinib in combination with carboplatin and paclitaxel, 68 patients were enrolled and median time to progression and overall median survivals were 6.1 and 16.9 months, respectively, for platinum-resistant patients, and 9.2 and 25.7 months, respectively, for patients with platinumsensitive disease. This study did report two secondary myelodysplatic syndromes and one secondary acute leukemia during treatment [94].

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Another small molecule tyrosine kinase EGFR reversible inhibitor is erlotinib. In a phase 2 study of recently diagnosed advanced stage ovarian cancer patients, patients received carboplatin and paclitaxel every 3 weeks up to six cycles plus daily oral erlotinib. EGFR amplification, which had been previously reported to occur in about 20% of ovarian cancer patients [95], was present in 15% of the evaluated tumors. The additional of erlotinib did not improve the pathologic complete response rate in this study compared to historical controls [96]. In a phase 3 trial-evaluating erlotinib as a maintenance therapy that included 835 patients and a median 51-month follow-up, erlotinib did not improve progression free or OS and no subgroup could be identified that might benefit from erlotinib maintenance [97]. In the recurrent setting, erlotinib in combination with bevacizumab [98], carboplatin and docetaxel [99], carboplatin and paclitaxel [100], and topotecan [101] have been evaluated with limited success. Cetuximab is a monoclonal antibody that targets EGFR and has been approved by the FDA for the treatment of advanced colon and head and neck cancers. Preclinical work suggested that cetuximab had synergy with platinum-based chemotherapies [102]. In a phase 2 clinical trial of cetuximab with carboplatin and paclitaxel, patients received IV cetuximab initially followed by weekly IV infusions. Carboplatin and paclitaxel were administered every 21 days for six cycles. Patients who received a complete clinical response continued to receive cetuximab for 6 months until toxicity or progression. Although tolerable as a primary therapy, there was no prolongation of PFS [103]. In the recurrent setting, patients received cetuximab as a monotherapy with 1 and 9 of 25 patients achieving a partial response and stable disease, respectively. Median PFS was 2.1 months [104]. In another phase 2 study evaluating recurrent platinum-sensitive patients, patient were retreated with carboplatin and cetuximab, but the response rate did not meet preset criteria for opening a second stage of accrual [105]. Trastuzumab and pertuzumab are recombinant, humanized monoclonal antibodies that are directed against HER2 and inhibit ligand-activated heterodimerization with other HER receptors. Lapatinib is a small molecular dual tyrosine kinase inhibitor of HER2 and EGFR. These are potential targeted therapy treatment options in women with HER2 amplification and overexpression, such as the success seen with the treatment of HER2/neu-positive breast cancer patients. Trastuzumab has been evaluated in the recurrent setting and eligible patients included those with HER2 overexpression as documented by immunohistochemistry. Of the 837 tumor samples that were screened, only 11.4% had the required expression level, and of those patients who were treated with IV trastuzumab, median progression-free

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interval was only 2 months [106]. In the platinumsensitive setting, carboplatin and paclitaxel were tested with and without pertuzumab. There was no significant difference between either arms in median PFS or response rates [107]. Pertuzumab has been evaluated in the recurrent platinum-resistant setting, and the objective response rate was 13.8% and 4.6% in patients who received gemcitabine and trastuzumab or gemcitabine and placebo, respectively [108]. Lapatinib is an orally available tyrosine kinase inhibitor of EGFR and HER2 by binding to the ATP-binding domain. Lapatinib monotherapy treatment in patients with recurrent disease has shown limited clinical efficacy [109], and combination lapatinib with carboplatin was associated with non-doselimiting toxicities [110]. Limited efficacy was also observed in combination lapatinib plus topotecan [111]. While these therapies have had limited successes, patients with mucinous ovarian cancer, which are known to have HER2 amplification and overexpression, may benefit from HER2 therapies. However, due the relative rarity of this subtype of epithelial ovarian cancer, accrual to a trial with only mucinous tumors would be difficult [112,113]. Seribantumab is a monoclonal antibody that binds to HER3, which then blocks heregulin-mediated (HRG) ErbB-3 signaling and then induces downregulation of the ErbB-3 receptor. A randomized phase 2 trial evaluated seribantumab in combination with weekly paclitaxel compared with paclitaxel alone in patients with platinum-resistant ovarian cancer. Of the 223 patients, median PFS was 3.75 months with seribantumab plus paclitaxel compared with 3.68 months with paclitaxel alone (P ¼ 0.864) [114].

FOLATE RECEPTOR-ALPHA Folate receptor-alpha, which is highly expressed on more than 90% of ovarian cancer cells [115], binds folic acid and transports folate into the cells by receptormediated endocytosis. This receptor allows cells to grow in microenvironments where there is a lack of folate, which provides a growth advantage to tumor cells [116,117]. Farletuzumab is a monoclonal antibody to folate receptor-alpha that leads to cell-mediated cytotoxicity, complement-dependent killing, and inhibition of growth under limited folate conditions [118,119]. In a phase 2 trial in patients with platinum-sensitive ovarian cancer, patients received single-agent farletuzumab or farletuzumab combined with carboplatin and paclitaxel (or docetaxel), followed by farletuzumab maintenance until progression. Of the 47 patients who received farletuzumab, 80.9% had normalization of their CA125, and a complete or partial objective response rate

was achieved in 75% with combination therapy [120]. In the subsequent phase 3 study evaluating 1100 women, two doses of farletuzumab were evaluated, and neither farletuzumab group had any significant difference in PFS from the placebo group [121]. In the PRECEDENT trial, vintafolide, a folic acid desacetylvinblastine conjugate, that binds to the folate receptor was evaluated in combination with pegylated liposomal doxorubicin compared to liposomal doxorubicin alone. Vintafolide is designed to use the internalization of the folate receptor to deliver the high-toxic desacetylvinblastine selectively to tumor cells. In this study, a folate receptor imaging agent, (99m)Tcetarfolatide [122], was used to select patients who might best benefit from treatment. Of the 149 patients evaluated, median PFS was 5 and 2.7 months, favoring the vintafolide plus pegylated liposomal doxorubicin arm (95% CI: 0.41
0.96; P ¼ 0.031). Patients who had between 10% and 90% folate receptor positive disease derived at least some improvement in PFS (HR, 0.873), but patients who completely lacked receptor expression received no benefit [123]. Unfortunately, the subsequent phase 3 trial, which compared vintafolide and pegylated liposomal doxorubicin (PROCEED), was terminated early due to an interim analysis which failed to show benefit (NCT01170650) [124].

P53 The most common alteration in high-grade serous ovarian cancers is mutations in P53 [5,125]. Although previously thought to be “undruggable,” recent agents have been developed that target P53. Cell cycle check points allow normal cells to repair their DNA prior to replication and division. Wee1 tyrosine kinase is involved in the phosphorylation that inactivates cyclin-dependent kinases 1 and 2 (Cdk1 and Cdk2) [126,127]. p53 mediates the G1 cell cycle checkpoint. Thus, cells with nonfunctional p53 become dependent on the G2 checkpoint. Wee-1-dependent phosphorylation of Cdk1 leads to activation of the G2 checkpoint and leads to a cell’s inability to repair its DNA. AZD1775 (MK-1775) is an inhibitor of Wee-1 kinase, and 25 patients were enrolled in a phase 1 study of refractory solid tumors. A partial response occurred in an ovarian cancer patient that also had a BRCA mutation. When combined with carboplatin in TP53-mutated ovarian cancer patients who were refractory or resistant (85%. Akt contains an N-terminal PH domain which binds to PI3K, a C-terminal rich in serine and threonine that is phosphorylated for kinase activation, and a central kinase domain. Akt is phosphorylated by phosphoinositide-dependent kinase 1 (PDK-1), integrated linked kinase (ILK), and MAP kinase-associated protein kinase-2 (MAPKAPK2) [78–80], but is negatively regulated by SH2 domain-containing inositol phosphatase (SHIP) [81,82] and phosphatase and tension homolog (PTEN) [83,84], which are known tumor suppressor genes mostly mutated/downregulated in cancers, including OVCA.

PI3-K/Akt Pathway in Ovarian Cancer Chemoresistance The PI3-K/Akt signaling pathway is critical in the regulation of OVCA cisplatin sensitivity. Akt is the most extensively studied downstream protein target of PI3-K that is implicated in OVCA tumorigenesis and chemoresistance. It is overexpressed, activated, and functionally altered at the DNA or mRNA levels in primary OVCA [32]. We have demonstrated that OVCA cells overexpressing a constitutively active Akt or amplified AKT2 are

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resistant to cisplatin-induced apoptosis compared to cells expressing low Akt levels [85]. Our laboratory together with others have demonstrated that the PI3-K/Akt pathway interact with other proteins such as HKII, XIAP, gelsolin, and p53 to regulate OVCA chemosensitivity. This implies that the mechanism of ovarian chemoresistance is multifactorial; hence, a holistic approach should be considered in its strategic management.

The Interaction of Akt Signaling With XIAP, p53, HKII, and Gelsolin Following the treatment of cultured cisplatin-sensitive OVCA cells (A2780s) with cisplatin, a decrease in XIAP content associated with increased procaspases-3 and -9 activation was observed [63]. Akt was cleaved when the whole-cell lysates were incubated with human recombinant active caspase-3. However, Akt cleavage was blocked by a specific caspase-3 inhibitor (DEVD). Also, XIAP downregulation induced Akt cleavage and apoptosis and these two responses were attenuated by the presence of DEVD [63]. These observations suggest that caspase-3 execute apoptosis through modulation of the antiapoptotic effects of Akt. On the other hand, overexpressing XIAP in A2780s OVCA cells resulted in increased Akt phosphorylation and decreased cisplatin-induced apoptosis, thus conferring chemoresistance [64,86]. This observation suggested that XIAP exerts its antiapoptotic effects through Akt activation but not its upregulation since total Akt remained unchanged. PI3-K inhibitor LY294002 was found to attenuate XIAP-induced Akt phosphorylation but increased cisplatin-induced apoptosis in A2780s that has wild-type p53 [63]. This effect was not seen in cisplatin-resistant OVCA cells (A2780cp) that harbor mutant p53, suggesting that p53 might also be a key player in determining OVCA chemoresistance [63]. Our previous study demonstrates that XIAP downregulation increases caspase-mediated MDM2 cleavage, p53 stabilization, and accumulation, which ultimately increased apoptosis in cisplatin-resistant cells (C13*) with wild-type p53 [65]. However, this effect is attenuated in A2780cp with mutant p53 after XIAP downregulation. Reconstituting A2780cp cells with wild-type p53 rendered the cells cisplatin-sensitive after XIAP downregulation, suggesting that the status of p53 is critical in determining XIAP-mediated chemoresistance [65]. Adenoviral vector containing a dominant negative Akt (DN-Akt) downregulates Akt activity in C13* and sensitizes the cells to cisplatin-induced apoptosis. However, such kind of response is not evident in A2780cp. Upon expression of wild-type p53 in A2780cp cells (mutant p53), downregulation of Akt and sensitization of cancer cells to cisplatin-induced apoptosis could be observed,

thus indicating that Akt-mediated chemoresistance is dependent on its ability to modulate p53 content and function [87]. There is also evidence that cisplatin induces p53 phosphorylation at the Ser15 sites through activation of ERK1 and ERK2 would result in increased apoptosis. Mutation of the Ser15 sites attenuates cisplatin-induced phosphorylation, leading to declined p53-mediated apoptosis [88–90].

TUMOR MICROENVIRONMENT AND CHEMORSISTANCE There are emerging evidences to support the importance of TME in cancer initiation and progression including chemoresistance in OVCAs. The TME entails a plethora of interactive cells that communicate with each other via cell-cell contact or the soluble factors that modulate the behavior of the tumors. Immune cells have been shown to work synergistically with chemotherapy to promote cancer cell death through improved antigen presentation and increased immune cell infiltration. However, the antitumor functions of immune cells could be downregulated or reversed to favor the growth of cancer cells and ultimately become resistant to chemotherapy treatment. Both innate (natural killer cells, macrophages) and adaptive (T cells and B cells) arms of the immune system have been implicated in OVCA chemoresistance. Endothelial cells play a key role in leukocyte extravasation and normal angiogenesis. The altered function of the endothelial cells result in the impairment of immune cells recruitment in the TME, decrease vascular adhesion molecule 1 (VCAM1), and promotes abnormal angiogenesis leading to tumor growth and chemoresistance [91]. Dysregulated endothelial cells may increase the secretion of immune-suppressive molecules such as prostaglandin E2 (PGE2) and Fas ligand (FASL), which attenuate the cytotoxic functions of T cells [91,92]. Cancer-associated adipocytes (CAAs) and fibroblasts (CAFs) are considered to be one of the most predominant cells in the TME contributing to the structural framework and homeostasis maintenance [91]. These cells when modulated by the tumor could secrete cancer promoting cytokines (TGF-β and VEGF) and chemokines that inhibit the cytotoxic functions of immune cells, thus facilitating the growth of tumor and promotion of metastasis by disrupting extracellular matrix (ECM)[91,93]. ECM are produced by cells and woven into a network of fabric in the TME. They provide structural support and regulate other cellular activities as well. During tumorigenesis, the tumor-associated ECM serves as a gateway for cancer cell invasion and proliferation. The interaction between cancer cells and ECM is crucial in ovarian carcinogenesis and chemoresistance [91].

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TUMOR MICROENVIRONMENT AND CHEMORSISTANCE

The vasculature system in the TME comprises the new formed vessels and the remodeling of existing mature vessels, both provide nutrients and oxygen to tumor cells to promote growth, invasion, and metastasis. The secretion of proangiogenic factors such as vascular endothelial growth factor (VEGF) by cancer cells, tumor-associated macrophages (TAMs), and cancer-associated fibroblasts (CAFs) increases tumor vascularization, thereby worsening patient outcome [91,94]. Tumor infiltrated lymphocytes (TILs) are immune cells that come from the circulation and presented in the tumor stroma as well as parenchyma. They consist of the members from both innate (macrophages and natural killer cells) and adaptive (T and B cells) immune systems. Although the normal physiological function of these immune cells should be to recognize and kill tumor cells, they are usually “re-engineered” to help tumor development instead [91]. In a functional immune system, tumor antigens should be sampled by antigen-presenting cells (APCs), such as dendritic cell (DCs), and macrophages then processed and presented to T cells to activate cytotoxic T cells for eradication of the tumor cells. This is known in the TME as the elimination phase [95]. The antigens could also be presented to helper T cells to get activated for proinflammatory and immune-stimulatory cytokines secretion that potentiates immune response. Tumor cells in their quest to combat the immune response may secrete inhibitory cytokines (TGF-β, IL-10) and chemokine ligand 2 (CCL2) to downregulate the cytotoxic functions of immune cells. This counter-attack mounted by the tumor cells are in balance with antitumor attack hence creating an equilibrium phase [95]. Although most tumor cells will be killed by the immune cells, some of the antigens in the tumor cells are mutated hence acquire resistance to tumor killing. This period of evading immune attack is referred to as the escape phase [95]. At this phase, the tumor becomes resistant, malignant, and metastasizes easily. Tumor-associated macrophages (TAMs) are phagocytic immune cells derived from monocytes that could either stimulate antitumor function or promote tumor growth depending on the local signal networks to which they are exposed. Circulating monocytes could be committed to either M1 or M2 macrophages depending on the cytokine stimulation [96]. Upon lipopolysaccharide (LPS) and/or interferon gamma (IFN-γ) stimulation, monocytes could be differentiated into M1 macrophages that are responsible for high-level IL-12 and IL-23 secretion but low levels for IL-10 [96]. On the other hand, when monocytes are stimulated by IL-4, they are differentiated into M2 macrophages, which secretes high-level IL-10, TGF-β, and IL-4 but low-level IL-12 [96]. TAMs are mostly M2 macrophage populations that secrete immunosuppressive cytokines to promote tumor growth, invasion, angiogenesis, metastasis, and chemoresistance

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[34,91,96]. The immune-suppressive role of TAMs has been reported in different types of cancers including OVCA, where increased TAM population is associated with poorer patient outcome, tumor aggressiveness, and chemoresistance [97]. Tumor-Infiltrating APCs are populations of immune cells that are supposed to recognize, process, and present tumor antigens to T cells to get them activated. Dendritic cells (DCs) are considered the most effective APCs. Antigen presentation is done via expression of costimulatory membrane ligands (CD40, CD80, and CD86) as well as secretion of cytokines (IL-12 and IFNs) [34,91]. In the TME, DCs are mostly downregulated through decreased infiltration and downregulation of costimulatory membrane ligands and IL-12 as well as IFNs, thereby attenuating the activation of T cells [98]. Reprogramming DCs to effectively recognize tumor antigens could be an effective way of boosting immune response toward OVCAs. Myeloid-derived suppressor cells (MDSCs) possess strong immune-suppressive effects against T cells, macrophages, DCs, and NK cells. Increased MDSC infiltration has been reported in OVCA as well as other type of cancers and is associated with poor patient outcome and chemoresistance [34,99]. MDSCs are effective in the production of indoleamine-2,3-dioxygenase (IDO), arginase 1, nitric oxide, and reactive oxygen species (ROS) known to inhibit the antitumor functions of T cells [34,91]. Targeting MDSCs in OVCA patients could contribute to eliciting strong antitumor responses. Metabolic activities in the TME contribute immensely to their fate of OVCA cells. These include amino acid synthesis, glycolysis, and de novo fatty acid synthesis, etc. Cancer cells and immune cells compete with each other for the limited nutrient resources such as glucose and amino acids in the TME for survival [34]. Cancer cells deprive immune cells from glucose and amino acids, which are otherwise needed by T cells and other immune cells for activation and differentiation. In addition to cancer cells, TAMs and MDSCs express arginase 1 and IDO, which deplete arginine and tryptophan, respectively, in the TME, thus decreasing their availability for T cell activation [91]. Kynurenine, a metabolite of tryptophan metabolism, also induces T cell dysfunction and promotes T-reg cells’ (immunosuppressive T cells) differentiation [34,100]. In addition to nutrient deprivation and metabolites accumulation, the local build-up of fatty acids is also detrimental to T cell activity and may enhance tumor growth. Natural Killer Cells (NK cells) are considered as innate cytotoxic lymphocytes that provide immediate response to viral infection and tumor formation. Unlike T cells that recognize tumor antigens on major histocompatibility complex (MHC), NK cells recognize tumor antigens in the absence of MHC thus allowing much faster immune response to contain a viral infection or tumor formation

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while other immune system members take time to produce tumor-specific cytotoxic T cells [101]. NK cells directly induce cell death via apoptosis or osmotic cell lysis. They express both activating (natural cytotoxicity receptor, NCR; ly49; CD16; CD94:NKG2) and inhibitory receptors (killer-cell immunoglobulin-like receptor, KIR; leukocyte immunoglobulin-like receptor, LIR; ly49) and depending on the stimuli received, could kill or inhibit killing [101]. They express and secrete perforin, IFN-γ, and proteases (granzymes) in their cytoplasm. Perforin creates pores on the membrane of target cells allowing granzymes and other secretory factors to enter and to induce apoptosis or cell lysis [91,102,103]. Aside from apoptosis, NK cells also involve antibody-dependent cell-mediated cytotoxicity (ADCC) and tumor cell surveillance. B cells or B-lymphocytes are populations of immune cells in the adaptive arm of the immune system that secrete antibodies upon activation and could also function effectively as professional APCs. Activated B-cells secrete inflammatory cytokines that potentiate the effects of T cells, macrophages, and NK cells to eliminate tumor cells [103]. Although the function of B-cells in tumor formation is not clear, studies have shown that increased CD20+ B-cells infiltration is correlated with improved outcome of OVCA patients [103–106]. Also, CD20 + B-cells colocalization with activated CD8+ TILs was associated with increased patient survival compared to CD8 + TIL alone [105,106]. Such effect may be due to the fact that B-cells could secrete cytokines that enhance CD8 + TIL activation and function. Although CD20 + B-cells are correlated with enhanced patient survival, CD19 + B cells, on the other hand, are predictive for poor survival [104]. This indicates that type of B cells in the TME is key to generate a strong immune response.

EFFECT OF CHEMOTHERAPY ON TILs AND IMPLICATIONS FOR IMMUNOTHERAPY Chemotherapy induces immunologic changes that augment antitumor immunity. Neo-adjuvant chemotherapy has been shown to be associated with increased infiltration or densities of CD3 +, CD8 +, CD8+ TIA-1 +, CD20 +, and PD-1 + TILs in HGSC [107]. However, CD68+ macrophages, MHC class 1 on tumor cells, FoxP3 + PD-1 + cells, IDO-1 + cells, and PD-L1+ cells (on both macrophages and tumor cells) remained unchanged [107]. Thus, neoadjuvant chemotherapy is associated with three response patterns: (i) TILhigh upregulated multiple immune markers after chemotherapy treatment, (ii) tumors with low TILs exhibited similar changes like the first group, and (iii) tumor with negative TILs remained negative. This suggests that chemotherapy

enhances preexisting TIL responses but fails to suppress immune-suppressive mechanisms [107]. Recent studies have also revealed that in OVCA, PD-L1 is highly expressed by CD-68 + macrophages rather than the tumor cells [108]. PD-L1 + macrophages frequently colocalize with both cytolytic (CD8 +, CD4+, granzyme B, and IFN-γ) and suppressive (PD-1 + TIL, CD25+ FoxP3 + Tregs, IDO-1, LAG3, and CTLA-4) subsets, resulting in a net positive association with survival [108]. The coexistence of cytolytic and suppressive subsets creates an immunological stalemate. Thus, explaining why multipronged approaches to immunotherapy are needed and should be tailored to the baseline features of the TME.

CROSS-TALK MOLECULES IN THE TUMOR MICROENVIRONMENT Intercellular communication is a fundamental process not only in the TME but also in the metastatic dissemination of tumor cells and chemoresistance. Cross-talk between cancer cells and neighboring cells could be mediated through several mechanisms, namely: (1) direct cell-cell contact, (2) cytokine/receptor interaction, and (3) vesicle-mediated interaction. As to whether these mechanisms act independently or dependently still remain a puzzle to be solved. Nevertheless, with the exception of the vesicle-mediated mechanism, the others have been extensively studied especially in the tumor microenvironment.

Tumor-Derived Extracellular Vesicles A plethora of extracellular vesicles are released by tumor cells into the TME which modulate neighboring target cells to favor tumor growth, angiogenesis and chemoresistance. These tumor cell secreted vesicles are heterogeneous and may include exosomes, microparticles, and apoptotic bodies. Regardless of the fact that these vesicles share some common features, they differ by their sizes, mechanism of formation, characteristic structural and molecular features, and compositions (proteins, nucleic acids and lipids, etc.). These distinctive features provide information about the cell of origin as well. Amongst these vesicles, exosomes and microparticles are the most studied in the context of tumor growth and chemoresistance. Exosomes are small vesicles measuring from 50 to 100 nm in diameter formed within endosomes by membrane invagination leading to the formation of multivesicular bodies (MVBs) [109]. Upon fusion with the plasma membrane, the exosomes are released into the extracellular space. Proteins such as ALG-2-interacting protein  2 (Alix2), endosomal sorting complex required for

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CROSS-TALK MOLECULES IN THE TUMOR MICROENVIRONMENT

transport (ESCRT), tumor susceptibility gene 101 (Tsg101), CD63, heat shock proteins (Hsps), and major histocompatibility complex II (MHC class II) are highly contained in exosomes hence used as markers for detection after sedementing at 100,000 g from appropriate samples (culture media and biological samples) [110]. Microparticles (MPs) are 100–1000 nm in diameter and formed by the outward blebbing of plasma membrane mostly under stress conditions [111]. MPs sediments at 20,000 g and is positive for Annexin V and other cell-specific surface markers. MPs, just like exosomes, shuttle proteins, lipids and nucleic acids that are involved in regulating tumor proliferation, invasion, and chemoresistance [112].

Cancer-Derived Extracellular Vesicles and Chemoresistance EVs-mediated drug resistance is frequently classified into three mechanisms: Neutralization of antibody-based drugs and export of small molecular drugs as well as transfer of biologically active materials (proteins, lipids, and miRNAs) [113–116]. Exosomes released from chemoresistant OVCA cells contained approximately threefolds more cisplatin compared to the exosomes from their sensitive counterparts, suggesting that cancer cells utilize exosome transport system for drug export [117]. It has also been demonstrated that cisplatin-resistant OVCA cells release exosomes that contain biologically active miR-21-3p which targets the neuron navigator 3 gene (NAV3) in the chemosensitive cells rendering them resistant to cisplatin treatment [118]. In functional studies, miR-21 was observed to be carried in exosomes released from CAAs or CAFs to OVCA cells. These functional exosomes suppressed paclitaxel-induced apoptosis in OVCA through their direct interaction with Apaf1 [119]. EVs derived from cisplatin resistant OVCA cells contain mutated SMAD4 that confers resistance of naïve A2780 (cisplatin-sensitive) recipient cells and is also associated with epithelial to mesenchymal transition (EMT) [120]. The efficacy of most immunotherapy has been compromized due to the ability of cancer-derived exosomes to bind to then inhibit therapeutic antibodies such as trastuzumab and rituximab. Cancer-derived EVs have also been shown to downregulate the cytotoxic functions of immune cells, thus enabling cancer cells to evade antitumor response. The underlying mechanism behind the transfer of EVs and selective uptake by immune cells are still largely unknown. Understanding how cancer-derived EVs modulate immune cells to facilitate tumor escape is a key to develop novel cancer therapies. EVs derived from malignant ascites of OVCA patients when internalized by monocytes induce protumorigenic cytokines as well as

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trigger TLR-dependent signaling pathways resulting in immunosuppressive mechanisms for cancer progression and chemoresistance [121]. Epithelial OVCAs (EOCs)derived exosomes contain miR-222-3p that when internalized by monocytes results in their polarization to M2 macrophages. These macrophages could also be differentiated to TAM-like phenotypes favoring tumor growth and chemoresistance [122]. The antitumor functions of NK cells are inhibited by exosomes released by OVCA cells through their interaction with the natural killer G2D (NKG2D) and CD226/DNAM-1-polio virus receptor/nectin-2 pathway [123]. Once the killing properties of NK cells are inhibited, the OVCA cells begin to dominate in the TME, undergo selective genetic mutations, and become resistant to chemotherapy treatment. Ascites-derived exosomes also contain FasL that induces apoptosis of activated CD8 + TILs [124]. Aside inducing T cell apoptosis, microparticles can downregulate the expression of TcR-associated ζ chain as well as Janus kinase 3 (JAK 3), thus attenuating T cell signaling cascade [124].

Plasma Gelsolin and Chemoresistance pGSN, a secretory/soluble form of gelsolin, differs from the cGSN by a N-terminal 24-amino acid signaling peptide and a cysteine residues disulfide bond between positions 188 and 201 designated as the “plasma extension” signal [36,37]. The major sites for pGSN synthesis are the skeletal, cardiac, and smooth muscles instead of the liver cells, which is a major organ that contributes most circulating proteins. pGSN is the fourth most abundant protein in the blood with an average concentration at 200–300 μg/mL in mammals [36,37,125]. pGSN is considered to be an extracellular actin scavenger to prevent the animal from further life-threatening conditions [39,126]. Such risk may present in an array of clinical conditions such as acute respiratory distress syndrome, sepsis, major trauma, malaria, and liver injury [39]. Elevated pGSN levels in the blood of colorectal, head-andneck, and OVCA patients are significantly associated with chemoresistance [127]. However, the mechanism behind this phenomenon has not been explored. Although pGSN is distributed by body fluids [125], little is known about its expression, secretion, and interaction with immune cells in the tumor microenvironment. Despite the fact that pGSN has been shown to interact with cell surface proteins such as integrin [128], detail interactions and resulting effects are yet to be studied. To date, whether pGSN interaction with integrin affects OVCA responsiveness to CDDP, and if this affects the functions of immune cells in the TME is not known, since most of the published data focused on average levels of pGSN in the blood, CSF, and inflammatory disorders.

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Cytokines/Chemokines and Chemoresistance

Immune Checkpoint Blockers

Cytokines are secreted molecules with diverse categories that play a pivotal role in normal physiologic and pathologic conditions. This collectively includes but not limited to chemokines, adipokines, pro/antiangiogenic factors, growth factors, proteases, and soluble receptors [129]. The aberrant expression and secretion of cytokines result in abnormal cytokine signaling may contribute to tumor growth, angiogenesis, metastasis, and apoptosis [129]. Emerging evidence suggest that cytokines secreted by cancer cells and stroma modulate the responsiveness of cancer cells to chemotherapy. Although the mechanism behind cytokine-mediated chemoresistance is not well established, they present as potential targets for treatment and diagnosis. Monitoring the circulatory levels of cytokines in the blood could be utilized as diagnostic biomarkers to predict chemoresistance, especially in OVCA where chemoresistance is a great obstacle to therapeutic success. C-C motif chemokine ligand 22 (CCL22) produced by tumor cells and TAMs serve as chemo-attractants to suppressive T regs in the tumor site, thus inhibit the antitumor response and thereby contribute to chemoresistance [130]. Increased C-X-C motif chemokine ligand 12 (CXCL12) secretion is associated with tumor growth and reduced survival in mice model [130]. Silencing CXCL12 gene expression or blocking its associated receptor, C-X-C motif chemokine receptor 4 (CXCR4), resulted in increased survival and decreased Treg recruitment [130,131]. In ovarian and breast cancers, increased levels of IL-6 are associated with chemoresistance and poor prognosis despite its poorly understood mechanisms [132–134]. Increased levels of VEGF, IL-10, and TGF-β in epithelial ovarian cancer (EOC) are positively correlated with Treg differentiation and immature dendritic cells infiltration, which could contribute immensely to drug resistance [135,136]. OVCA patients in the advanced stages have increased levels of IL-6, IL-8, IL-10, and TNFα in their blood [132–134]. As to whether these cytokine elevations have a role in chemoresistance is yet to be explored.

Cancer immunotherapy has several recent breakthroughs and hence gained significant attention from both cancer research community and clinical therapeutics providers. Being recognized as a robust immune regulatory mechanism, the programmed death 1 (PD-1) pathway has warranted its use as prognostic marker and therapeutic candidate in the treatment of cancer. PD-1 is constitutively expressed as a surface receptor by activated CD4 and CD8 T cells as well as other immune cells such as NK cells [137]. PD-1 has two major ligands, the PD-L1 and PD-L2, that are expressed by tumor cells, tumor-associated macrophages (TAMs), dendritic cells, T cells, B cells, and myeloid-derived suppressor cells (MDSCs) [137]. The interaction of PD-1 and PD-L1/2 results in reduced T cell activation, proliferation, differentiation, cytokine production, tolerance, and increased apoptosis [138]. Blocking the PD-1/PD-L1 interaction with monoclonal antibodies rescues T cell antitumor function resulting in a more potent immune response [138,139]. In a phase I trial that 17 OVCA patients were recruited, PD-L1 blocking antibody (BMS-936559: Bristol-Myers Squibb) achieved 1 partial response (PR) and disease stabilization (SD) in 2 patients [140,141]. Avelumab (anti-PDL1 antibody) resulted in 4 PR and 11 SD in a phase I trial (NCT01772004) [140]. Nivolumab (anti-PD1 antibody) achieved 2 complete responses (CRs), 2 PR, and 6 SD in a phase II trial (UMIN000005714) in which 20 OVCA patients were recruited [139]. Another PD-1 blocking antibody (Pembrolizumab), achieved 1 CR and 2 PR amongst 26 patients in a phase I trial (NCT02054806) [140]. Cytotoxic T lymphocyte-associated protein 4 (CTLA4) is an inhibitory receptor expressed on T cells after activation. CTLA4 inhibits T-cell activation, proliferation and antitumor activities after binding to CD80 or CD86 on APC [142]. CTLA4 is also constitutively expressed on suppressive T regs. Blocking the interaction between CTLA4 and CD80/86 reverses its immune-suppressive effect and potentiates antitumor effects [142]. Ipilimumab, a blocking antibody against CTLA4, is approved for the treatment of metastatic melanoma is also investigated in other solid tumors [143]. Currently, patients with recurrent platinum sensitive OVCA have been recruited (NCT01611558) into a phase II trial with results pending. Tremelimumab, another blocking antibody against CTLA4, is being tested in patients with solid tumors in a phase I trial [140]. Although these immune checkpoint blockers demonstrated promising results in the treatment in OVCA, there are no concrete data about their efficacy in chemoresistant OVCA patients. As to whether chemoresistant OVCA patients will benefit from such novel treatment regimens are yet to be demonstrated. There is therefore

MODERN TREATMENT STRATEGIES FOR OVARIAN CHEMORESISTANCE Chemoresistance is a major setback in OVCA treatment. Although patients initially respond to the first-line treatment—surgical debulking and chemotherapy, a significant proportion recurred and became resistance to treatment. In view of this, major research works are underway to explore effective antichemoresistance strategies to achieve better therapeutic success.

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a need for advanced personalized treatments where chemoresistant patients would benefit from treatment based on the alterations in their tumor microenvironment.

Cell-Based Immunotherapies Advanced therapeutics have been proposed through reprogramming autologous immune cells such as T-cells, NK cells, and DCs ex vivo then reintroducing them into the patients to attack cancer cells. OVCA tumor-associated antigens (TAAs) such as NY-ESO-1, CA-125, and MAGE-A1 can effectively stimulate and activate T cells. Such properties of these antigens have been explored extensively and used as peptide vaccines [144]. DCs and T cells, when pulsed with OVCA TAAs, have shown promising results [144]. However, modest immunological responses were observed with peptide vaccines in recurrent OVCA patients. Adoptive cell transfer (ACT) therapy: This involves the ex vivo expansion of autologous specific TILs and reintroducing them into the same patient to generate effective antitumor response. Patients received TILs demonstrated enhanced overall survival and favorable clinical outcome [103]. ACT of TILs after surgical debulking and chemotherapy resulted in a significant PFS in a pilot trial involving 13 recurrent OVCA patients [103,145,146]. T cells could also be genetically engineered to express either tumor-specific T cell receptors (TcRs) or chimeric antigen receptors (CARs) [145]. Despite the promising results associated with engineered T cells, the associated toxicities (grades 3 and 4) are the main obstacles to their acceptance into further clinical practice [145]. Refined FRα CAR is currently being tested in advanced OVCA patients in a phase I trial [144]. DC-based vaccine: This treatment approach involves the isolation of autologous DCs and priming them with TAAs ex vivo in order to enhance DC maturation and antigen presentation. These primed DCs are then reinfused into the respective patients to initiate strong tumor-specific CD8 + T cells response [147]. In a phase I trial, where DCs were primed with either HER-2/ neu or MUC-1 peptides, antigen-specific CD8 + T cells were detected in 2 advanced OVCA patients [147,148]. HER-2/neu-based DC therapy, Lapuleucel-T (Neuvenge, Denderon), in a phase I trial resulted in a short-term disease stability in 2 out of 4 OVCA patients [147,148]. These interesting results suggest that priming DCs with specific tumor-associated antigens is promising and more detailed fine-tuning should be done to ensure maximum antitumor responses. Combining this treatment with conventional chemotherapy could increase patients’ survival and help tackle the issue of chemoresistance.

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Targeting Epigenetic Changes Epigenetic changes such as DNA methylation and histone acetylation could modulate immune responses in the OVCA TME, thereby favoring tumor growth and chemoresistance [149]. DNA-histone interactions downregulate the expression of MHC-II, which correlates with increased tumor growth [150]. Histone deacetylase inhibitor (HDACi) treatment, however, reverses this effect leading to increased MHC-II expression and enhanced immune response [150]. Although preclinical data of HDACi in OVCA showed significant results, phase II trials of belinostat (Spectrum) [151] and vorinostat [152] only showed modest response. Combination therapy with other treatment modalities may increase patient response and disease stability.

NATURAL FOOD COMPOUNDS Natural food compounds have emerged as promising therapeutics against chemoresistant OVCA. Food phytochemicals contain certain bioactive molecules from natural plants and are known to modulate key regulators of apoptosis. These functional phytochemicals have remarkable effects in inhibiting survival pathways involved in tumorigenesis and chemoresistance. Phytoalexin resveratrol (RSV) is a stilbene extracted from grapes and mulberry and known for possessing strong anticancer properties. RSV inhibits ovarian cancer growth and also sensitizes cancer cells to cisplatin. We demonstrated that RSV enhances CDDP sensitivity in OVCA through p53-mediated expression of NOXA, increased XIAP degradation via ubiquitin-proteasome pathway, and enhanced caspase-3 activation [153]. These effects were also associated with Drp1-dependent mitochondrial fission. RSV selectively inhibits glucose uptake and induce apoptosis regardless of p53 status in vitro [153]. We have also shown that diarylheptanoid hirsutenone extracted from tree back of Alnus hirsute var. sibirica, effectively induced CDDP sensitivity in ovarian and cervical cancer cell lines—C13*, A2780cp, Hey, OVCAR433, SKOV-3, and OCC-1- regardless of p53 status [154]. Hirsutenone treatment enhanced XIAP degradation via ubiquitin-proteasome pathway and facilitated the translocation of AIF from the mitochondria to the nucleus [154]. A yellow pigment from Curcuma longa called Curcumin is an effective inhibitor of the cell cycle and promoter of caspase-mediated apoptosis [155]. It also inhibits SERCA activity and disrupts calcium homeostasis, thereby inducing apoptosis in OVCA cells [155]. In addition, we have recently demonstrated that Saikosaponin-d (Ssd), a major triterpenoid saponin derived from Bupleurum falcatum L. (Umbelliferae), sensitizes chemoresistant OVCA cells to CDDP-induced

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apoptosis by facilitating mitochondrial fission and G2/M cell cycle arrest in vitro. Mechanistic studies indicate that Ssd increases cytosolic Ca2+ concentration and facilitates CDDP-induced mitochondrial membrane potential loss and mitochondrial fission in chemoresistant cells. Ssd also downregulates PPM1D content, resulting in the activation of Chk1, phosphorylation of Cdc25c and Cdk1 (Tsuyoshi H. et al. In press, Oncotarget, 2017). Although remarkable preclinical data have been demonstrated with natural food compounds, clinical trials and FDA acceptance are still some miles away.

NANOTECHNOLOGY/NANOMEDICINE Nanotechnology has gained significant attention as a potentially alternative approach to cancer treatment due to its unique properties in size, advanced delivery, precise molecular level control of cellular and TME interactions, as well as unique physical chemical properties for molecular imaging, biomarker sensing, integrated multimodal therapy and even theranostics (combination of diagnosis and therapy in one platform) [156]. Nanomedicine is also known to provide advantages of minimal toxicity and solutions for chemoresistance. Various nano-formulations have been developed in the past decades and some of these were FDA approved for clinical use. These include liposomes, micelle-like structures, polymer aggregations, as well as inorganic nanomaterials or organic-inorganic hybrid materials [156]. Gold nanoparticles (AuNPs) has been shown to inhibit OVCA growth and metastasis through its ability to reverse EMT, which otherwise confers resistance to cancer cells. AuNPs sensitizes chemoresistant OVCA cells to CDDPinduced death and also decreases the expression of key stem cell markers—MDR1, Sox2, CD133, ALDH1, CD44, and ABCG2 through downregulation of the Akt and NFkB signaling pathways [157]. Orthotopically implanted OVCA tumor was sensitized by AuNPs to CDDP resulting in a significant decrease in tumor growth [157]. Doxil, a PEGylated liposomal doxorubicin, has been approved by FDA for the treatment of OVCA (NCT00945139, NCT00862355) [156]. Liposomal topotecan (NCT00765973) and liposomal lurotecan (NCT0001017) are currently being tested in a phase I and II trials, respectively, in OVCA patients [156]. Xyotax, a paclitaxel poliglumex, is also under phase II trials in OVCA patients (NCT00060359) [156].

PERSONALIZED CANCER THERAPY: THE FUTURE? The underlying mechanisms of OVCA chemoresistance are multifactorial; hence, effective treatment should

encompass multiple therapeutic approaches. Targeting only one oncogene or signaling pathway does not provide significant tumor suppression due to the heterogeneity and multifaceted nature of the tumor in each patient. Most cancer treatments target either one of the following: tumor cell receptors, immune cells, activated pathways, mutations, and oncogenes. However, the major setbacks have been tumor heterogeneity, genetic instability, chemoresistance, and TME. This seems to explain why treating patients with the same diagnosis using the same therapy has not yielded significant therapeutic success. Most of the single-target therapies approved by FDA and those in clinical trials only offer prolonged survival for few cancer patients while the majority do not respond well. There is therefore the need to focus on individual patient-tailored treatment that gathers extensive cellular, molecular, and physical information of the patient to effectively design therapies [158]. This is termed personalized cancer therapy (PCT). PCT takes into consideration patient’s diagnostic history, molecular and genetic profiles of the TME, cancer biomarkers, and the patient. The bio-informatics data of the patient are usually applied to design effective therapeutic regimens with minimized toxicities. Humanized mouse models and 3D-organoid systems have been developed to study the interaction between patientderived cells and the stroma to identify potential dysregulated pathways contributing to drug resistance in a clinically mimic in vitro environment [159,160]. Using carefully selected molecular targets from a patient; the tumor could be better managed and treated while systemic toxicities could be minimized. Advanced technologies have made it easy to sequence patients’ tumor and normal cells to detect possible players involved in driving tumor growth and chemoresistance. Using such information together with patient-derived biomarkers and TME environment, precise cocktail therapies could be designed to effectively treat each patient according to the information gathered in an evidence-based manner. This could replace the “one-size-fits-all-therapies” in the clinic. Although this patient-tailored treatment approach seems probable, contingency health care plans should be considered to make treatment affordable to patients.

CONCLUSION Chemoresistance is a major problem in achieving longterm therapeutic success in OVCA patients. Although various key players such as oncogenes and tumor suppressor genes have been identified over the years as contributing factors to developing chemoresistance, there is currently no effective way to successfully and comprehensively solve these critical issues. Increasing evidence

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has also linked the influence of the TME to OVCA chemoresponsiveness to cisplatin. This supports the argument that, the underlying cause of chemoresistance is multifactorial, and hence will demand multidisciplinary treatment approaches. Novel therapies have been designed and currently in various phases of trials. However, only a hand full of patients responds while the majority of patients failed ultimately. This is irrespective of the fact that these patients have the same clinical initial diagnosis. This calls for individually tailored cancer therapeutics. OVCA patients should not be put in the same box during treatment but rather be treated according to the molecular and TME profiles detected in each individual. PCT explore the cellular and molecular dynamics of each patient’s tumor, and utilize such information to design a personalized therapy for each patient. This approach guarantees a holistic means of attacking chemoresistance and minimizing toxicities at the same time.

Glossary Chemoresistance—Most cancer cells are initially sensitive to chemotherapy-induced death; however, few populations develop mechanisms that render them resistant to chemotherapy. These resistant mechanisms may be due to genetic mutations and/or changes in the tumor microenvironment. The tumor microenvironment (TME) comprises of a plethora of soluble and cellular environment such as fibroblast, cytokines, immune cells and blood vessels that interact to modulate the growth of cancer cells. Extracellular vesicles (Evs) are released via exocytosis by multivesicular bodies or shed from the plasma membrane from all types of cells. Evs transport lipids, proteins and nucleic acids between cells and have a significant effect on the biological activities of the recipient cells. Immunotherapy involves harnessing the body’s immune system to fight cancer. This entails but not limited to monoclonal antibodies, immune-cell therapies, oncolytic virus therapy and cancer vaccines. Personalized cancer therapy takes into consideration the genetic make-up of the patient’s tumor to aid in the development of treatment strategies that are more effective but with lower side effect.

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Index

Note: Page numbers followed by f indicate figures and t indicate tables.

A

ABCA1, 88 Abdominal and total body irradiation, 503 Abnormal expression, in reproductive diseases, 213–214 Acetylation, 161 Acne, 419, 423 A2780cp cell lines, 576, 582 Acquisition of granulosa cells, 34–35 luteolytic capacity, 270–271, 272f ACTCH. See Adrenocorticotropic hormone (ACTH) Activin, 27, 35–37, 487 A, 100 abnormalities in PCOS, 101 α and β subunit synthesis, 97 autocrine/paracrine roles, 98 biological action, 97 cleavage of propeptide, 96 definition, 95–96 estrogen receptor regulation, 97–98 knocking out mice models, 98 molecular structure, 96 ovarian steroidogenesis, 115–116 signaling, 96–97 stimulates FSH production, 95–96 Activin-like kinase (ALK), 35 Activin receptor-type IIA, 304 Acute leukemia, 498 Adipokines, 522 Adiponectin, 448 A disintegrin and metalloproteinase 10 (ADAM-10), 28 Adoptive cell transfer (ACT) therapy, 587 Adrenal tumors, 416 Adrenocorticotropic hormone (ACTH), 416, 476 Adult granulosa cell tumors (AGCTs), 518 Advanced glycation end products (AGEs), 452–454, 454f Adverse effects, 487–488 AE-PCOS. See Androgen Excess and PCOS (AE-PCOS) AFC. See Antral follicle count (AFC) Aflibercept, 563 AGEs (advanced glycation end products), 452–454, 454f Aging dominant ovulatory follicle, 59 mitochondrial mtDNA mutations, 196

ovarian concept of, 380 inhibin B, 99 physiology of, 381–383 reproductive, 339 Ahch, 461 AKT interaction with XIAP, p53, HKII and gelsolin, 582 isoform-specific alterations of, 519 and PI3K signaling pathway aberrations within, 555 description, 581 downstream effector, 555 granulosa cell differentiation and proliferation by, 40 inhibitors, 556 and MEK inhibitor combination therapies, 565 in ovarian cancer chemoresistance, 581–582 primordial follicle activation, 29–32 Aldosterone, 476 Alisertib (MLN8237), 559 Alkylating agent dose (AAD), 501 Alnus hirsute var. sibirica, 587–588 American Society for Reproductive Medicine (ASRM), 384, 493–494 American Society of Clinical Oncology (ASCO), 493–494 AMH. See Anti-M€ ullerian hormone (AMH) AMP-activated protein kinase (AMPK), 242–243 activation, by PGF2α, 243–244 activation loop, 242–243 AMP:ATP/ADP:ATP ratios, 242–243 inactivation, 243 inhibition, by LH, 243 pharmacological activators, 242–243 phosphorylation, 243 to restore energy homeostasis, 242–243 Anastrozole, 557 Androgen Excess and PCOS (AE-PCOS), 417–418, 423 Androgen receptor (AR), 10, 442, 465 Androgen-resistant female mouse models (ARKO), 10 Androgens, 10, 41–42 Androgen-secreting tumors, 416 Androstenedione, 418 Angiogenesis and neovasculature, 522–524

595

Angiopoietin-2, 498 Anovulation chronic description, 419 infertility and, 421 diverse etiology, in women, 226–227 Anovulatory waves, 58–59 Anti-CTLA-4 antibodies, 561 Antigen presenting cells (APCs), 562f Antimesothelin antibodies, 559 Anti-M€ ullerian hormone (AMH), 29, 58, 61, 418, 421, 487, 503 age-specific, 383f assay, 409 in assisted reproductive technology age-specific normative values, 407 fertility preservation, 408–409 individualization, 407–408 polycystic ovarian syndrome, 408 prognostication counseling, 407–408 concentrations in women with PCOS, 119 in female reproduction, 112–113 FSH-induced aromatase expression, 116 FSH responsiveness inhibition, 404–405 functions, 403 in human ovary, 110 levels, factors influencing, 405–406, 406t limitations, 409 mutations, 403 overexpression of, 404 primordial follicle recruitment inhibition, 404 receptors, 403–404 role, 403–404 sex-dimorphic expression pattern, 403 signals, 403–404 stimulation, 406–407 TGFβ superfamily member, 403–404 Antral follicle count (AFC), 63, 348, 407, 503 Antral follicles age-related changes in, 59 FSH-induced growth, 8 preantral and early development, 52–53 recruitment of, 53–54 Apoptosis, 5–6, 299–300, 487–488 ARID1A mutations, 517, 548 ARIEL2, 551 ARIEL3, 551–552 ARIEL4, 551–552 Aromatase, 61 Aromatase mutations (CYP 19), 477

596 ART. See Assisted reproductive technology (ART) Aryl hydrocarbon receptor, 487–488 AS101, 502 Aspirin, 351 Assisted reproductive care, 486 Assisted reproductive technology (ART), 403 anti-M€ ullerian hormone age-specific normative values, 407 fertility preservation, 408–409 individualization, 407–408 polycystic ovarian syndrome, 408 prognostication cousenling, 407–408 definition, 380–381 inhibin B, 101 IVF treatment adverse outcomes, 383–384 age dependent, 384 multiple pregnancies, 383–384 offering centers, 384 for oldest patient population, 384–385 U.S. centers report, 384 U.S. live birth rate, 384, 384f procedures, 506 Ataxia telangiectasia and Rad3-related protein (ATR), 575–576 Ataxia telangiectasia mutated protein (ATM), 575–578 Atezolizumab, 562 ATP-binding cassette (ABC) pump inhibitor, 532 Atresia focal point of, 5 schematic representation of, 4f AURELIA phase 3 trial, 554 Aurora kinases, 559 Autoimmune disorders, 493 Autophagy, 487–488 Autophagy-related gene 7 (Atg7), 300 Autosomal-recessive disorders, 475t Avastin, 522 Avelumab, 562 Avon Longitudinal Study of Parents and Children (ALSPAC), 446–447 AZD1775 (MK-1775), 558–559

B

Balbiani body, 24–25 Basic helix-loop-helix (bHLH) transcription factor, 472 B cell lymphoma-2 (BCL-2) family, 25 B cells, 584 Beclin-1 (Becn1), 300 Belinostat, 560, 587 β-catenin, 298 Beta-cell function, 440 Bevacizumab, 553–555, 557, 561, 563–565 Binding proteins, 487 Binimetinib, 556 Bisphenol-A (BPA), 151–152, 444–446, 486–487 Blepharophimosis-ptosis-epicanthus (FOXL2), 479 Blimp1. See Positive regulatory domain 1 (Prdm1) B-lymphocytes, 584 BMP15. See Bone morphogenetic protein 15 (BMP15) Bone marrow transplantation, 493

INDEX

Bone morphogenetic protein 15 (BMP15), 160, 487 follicular growth controlled by, 35–36 mutation, 36, 118–119, 464 oocyte regulation of follicle selection, 11–13, 14f protein, 118–119 TGF-β superfamily members, 109, 118 Bone morphogenetic protein 4 (BMP4) pathway, 296, 298 Bone morphogenetic protein receptor type II (BMPRII), 35 Bone morphogenetic proteins (BMPs), 99, 107 Booroola (FecB), 35–36 BPA. See Bisphenol-A (BPA) BRAF mutations, 514, 516–517 BRCA mutations, 511, 518–519, 551–553, 558–559 BRCA1 mutations, 515–516 Breast carcinoma xenograft models, 539–540 BRG1, 548 Buparlisib, 565

C

CAAs. See Cancer-associated adipocytes (CAAs) Cabozantinib, 565 CAG repeat sequence, 442 cAMP response element-binding protein (CREB), 38–39 Cancer-associated adipocytes (CAAs), 521–522, 582 Cancer-associated fibroblasts (CAFs), 520, 582 Cancer cell reimplantation, risk of, 498 Cancer-derived extracellular vesicles, 585 Cancer Genome Atlas, 536–537 Cancer stem cell (CSC), 521, 538–539 Candidate gene studies, 439–441 beta-cell function, 440 obesity, 440 ovarian folliculogenesis, 440–441 steroid production and metabolism, 440 Carbohydrate-deficient glycoprotein (CDG), 476 Carboplatin, 547, 552–554, 556–560, 563–565 Cardiovascular disease (CVD), 422–423 CASPASE-2, 25–26 CCAAT/enhancer binding proteins alpha and beta (CEBPα/CEBpβ), 219–221 CCC. See Clear cell carcinoma (CCC) C-C motif chemokine ligand 22 (CCL22), 586 CDDP. See Cisplatin (cis-diamminedichloroplatinum (CDDP)) CDG (carbohydrate-deficient glycoprotein), 476 CD8(+) T cells, 520 Cediranib, 562–564 Cedirinib, 563–564 Cell-based immunotherapies, 587 Cell cycle molecules, nuclear maturation CDK1 activity, 166–168 CSF activity, 168 cyclin B1, 167–168 maturation-promoting factor, 166–167, 167f mitotgen-activated protein kinase, 166–167, 167f

Cell plasticity, 529, 535–536 Cellular cortex, 159 Cellular signals germ cell nest breakdown and primordial follicle formation, 24–29 growth factors and somatic cell cycle progression, 27–28 neonatal oocyte survival, molecular control of, 25–26 notch signaling, regulation, 28–29 steroid hormones, 26–27, 26f during preantral follicular development, 34–38, 34f follicular growth, control of, 35–36 granulosa cell proliferation, regulation of, 36–37 Notch signaling, 37–38 ovary, TGF-β signal transduction in, 35–37 theca cell recruitment and specification, molecular mechanisms, 38 during preantral follicular development-, 34–38, 34f Cerebellar ataxia, 473 Cetuximab, 557 CGA. See Common glycoprotein alpha (CGA) Chemoresistance, 538, 575–576, 577f cancer-derived extracellular vesicles, 585 cell-based immunotherapies, 587 cellular mechanisms, 576 and cisplatin action, mechanism of, 575–576 cross-talk molecules in TME, 584–586 cytokines/chemokines, 586 gelsolin (GSN), 576–577, 578–580f, 582 hexokinase II, 577–578, 582 immune checkpoint blockers, 586–587 immunotherapy, 584 inhibitor of apoptosis proteins, 578–581 nanotechnology/nanomedicine, 588 natural food compounds, 587–588 P53, 577–578, 582 personalized cancer therapy, 588 PI3-K/Akt pathway, 581–582 plasma gelsolin, 585 targeting epigenetic changes, 587 TILs, chemotherapy on, 584 tumor-derived extracellular vesicles, 584–585 tumor microenvironment, 582–584 Chemotherapeutic agents, 493 Chemotherapy, 493, 498, 511–512 Chemotherapy-related infertility risk, 501 Chinese-ancestry studies, 439 ChIPSeq, 530 Cholesterol acquisition, storage, and trafficking of, 83–84, 84f compartmentalization, 83 de novo synthesis of, 85 free, 86–87 gene expression, regulation of, 92 hypercholesterolemia, 85 intracellular trafficking, 87 lipid droplets, 86–87, 86f lipoprotein-mediated uptake high-density, 85 low-density, 84–85 for steroidogenesis, 85–86

INDEX

plasma membrane, 84 steroidogenic acute regulatory protein structure, 86–87, 86f transcriptional and post-translational mechanisms, 87–88 Cholesterol esterases, 86–87 Cholesterol side-chain cleavage enzyme (CYP11A1), 237–238 Chromatin immunoprecipitation (ChIP-seq), 171–172 Chronic anovulation, 419, 421 Cilia, 516 Cisplatin (cis-diamminedichloroplatinum (CDDP)), 560 FLIP, 577 GSN, 579–580f mechanism of, 575–576 response rate (RR), 576 Clear cell adenofibromas, 517 Clear cell carcinoma (CCC), 514, 515f, 517 Clomiphene, 425–426 Clomiphene citrate (CC), 425–426 CLPP, 473 Cluster of Differentiation (CD), 530 COC. See Cumulus-oocyte complex (COC) Coenzyme Q10 (CoQ10), 339 Color flow Doppler ultrasonography, 62 Combination therapy, OVCA, 562–565 and emerging antiangiogenic therapies, 563–565 PI3K/AKT and MEK inhibitor, 565 Combined oral contraceptives (COCs), 506 Common glycoprotein alpha (CGA) mutation, 132 polymorphisms, 132 structural features of, 128, 129f Connective tissue growth factor (CTGF), 27 Connexin-37 (Cx37), 304 Continuous Recruitment Theory, 53–54 Contraception, 227–228 hormonal, 59 mechanism of ovulation, 227–228 options, 504, 505t oral, 352, 354, 424 Contraceptives targeting ovulation, 227–228 Controlled ovarian hyperstimulation (COH), 380 Controlled ovarian stimulation (COS), 345 Corpus hemorrhagicum (CH), 62 Corpus luteum (CL). See also Luteolysis luteal rescue during early pregnancy, 284 IFNT, 285 maternal recognition period, 285 PAG, 285 pig, 283 primate, 281–282, 283f rodent, 282–283, 283f in ruminant, 279–281, 283f progesterone production extensive vascular network, 238 functional lifespan, 238 immune cell infiltration, 238 LH, 239 luteal regression, 238 menstrual or estrous cycle, 238

Corpus luteum (CL) formation hypoxia-inducible factor 1 activation, 256–257 HIF1A-dependent genes, 256–258, 256–257f independent regulation, 258–259 LH-dependent gene expression, 256, 257f mechanisms, 256, 256f microRNA-210, 259 luteal angiogenesis bovine ovary 1-day postovulation, 260, 260f FGF2, 260–261 PGE2, 261 VEGFA, 260–261 Cre Driver Mouse Lines, 297t Cre-loxP system, 28 cre recombinase-loxP system, 296 CSB-PGBD3 (POF11), 472 C-terminus cytoplasmic gelsolin (C-cGSN), 576–577 CTNNB1 mutations, 517 Cuboidal granulosa cells, 297f, 303 Cullin ring-finger ubiquitin E3 ligase-4 (CRL4) complex, 223 Cumulus cells (CCs), 4, 116–117 Cumulus granulosa cells, 165 Cumulus-oocyte complex (COC), 306 expansion, 217–218 activation, 220f, 221–222 extracellular matrix production, 222 granulosa cell EGF-like factors, 222–223 oocyte meiosis, resumption of, 221–222 ovulatory pathways, 223 functioning of, 13 OSF signaling, 14–15 release at ovulation time, 306 Cumulus-oophorus complex, formation and expansion, 116–117 Curcuma longa, 587–588 Curcumin, 587–588 Cushing’s syndrome, 416–417 C-X-C motif chemokine receptor 4 (CXCR4), 586 Cyclic adenosine monophosphate (cAMP), 16–17 Cyclic guanosine monophosphate (cGMP), 169 Cyclic Recruitment Theories, 53–54 Cyclin B1 cdc25, 167–168 levels, nuclear maturation, 184 translational control, 167–168 Wee1B, 167–168 Cyclin B (CYB), 160 Cyclin-dependent kinase (CDK), 560 Cyclin-dependent kinase 1 (CDK1), 160, 182, 305–306, 558–559 Cyclin-dependent kinase inhibitor, 27–28 Cyclooxygenase (COX) pathway, 306 Cyclophosphamide, 502, 552–553, 563–564 Cyclophosphamide equivalent dose (CED), 501 Cyp26b1, 36 Cysteine-aspartic proteases (caspases), 300 Cytochrome P450, 88

597 Cytochrome P450 side-chain cleavage CYP11A1, 245–246 FDX1, 245 FDXR, 245 steroidogenesis and, 90–91 Cytokines, 277–278 and chemoresistance, 586 CXCL8, 278 FAS-FAS ligand system, 277–278 IFNG, 277–278 immune cell infiltration and inflammatory mediators, 226 matrix remodeling, 278 production and regulation, 277 TNF, 277 transcription, 277 Cytoplasm molecular maturation BTG4 and CCR4-NOT RNA deadenylase, 190–192, 191f dormant mRNAs, translational activation of, 189–190 maternal mRNA decay, 190 ZAR1 and ZAR2, 192 zinc finger protein 36 like 2, 192 organelle maturation cortical granule migration, 193–194 cytoplasmic maturation, improvement of, 195–196 cytoplasmic quality, evaluation of, 195–196 cytoskeleton dynamics, 195 ER and Golgi complex, redistribution of, 194–195 mitochondrial number and distribution, 192–193, 194f Cytoplasmic gelsolin (cGSN), 576 Cytoplasmic lipid droplets, 86–87 Cytoplasmic maturation early embryonic development, 157–158 fertilization, 157–158 maternal effect genes, 158 reorganization, 158–159 Cytoplasmic molecular/organelle maturation, oocyte meiotic maturation BTG4 and CCR4-NOT RNA deadenylase, 190–192, 191f cortical granule migration, 193–194 cytoplasmic maturation, improvement of, 195–196 cytoplasmic quality, evaluation of, 195–196 cytoskeleton dynamics, 195 dormant mRNAs, translational activation of, 189–190 ER and Golgi complex, redistribution of, 194–195 maternal mRNA decay, 190 mitochondrial number and distribution, 192–193, 194f ZAR1 and ZAR2, 192 zinc finger protein 36 like 2, 192 Cytosine-thymidine transition (C566T), 477 Cytostatic factor (CSF), 168 Cytotoxic T lymphocyte antigen-4 (CTLA-4), 561

598 Cytotoxic T lymphocyte-associated protein 4 (CTLA4), 586

D

Dacarbazine, 502 Danusertib (PHA-739358), 559 DAPT, 28 Dasatinib, 560 DAZL, 475 DC-based vaccine, 587 DDE (dichlorodiphenyldichloroethane), 152 DDT (dichlorodiphenyltrichloroethane), 151–152 Death receptors, 577–578 DEET (N,N-diethyl-meta-toluamide), 151 DEHP. See Di(2-ethylhexyl) phthalate (DEHP) Dehydroepiandrosterone sulfate (DHEAS), 415 Delayed time-to-pregnancy (TTP), 486 Demcizumab, 559 Dendritic cells (DCs), 225, 583 DENND1A, 442 De novo methylation, 212–213 Denys–Drash syndrome, 474 Depot medroxyprogesterone acetate, 506 DEVD, 582 DHEAS, 428 DHT. See Dihydrotestosterone (DHT) Diarylheptanoid hirsutenone, 587–588 Dibutyl phthalate (DBP), 151–152 DICER1 mutations, 518 Dichlorodiphenyldichloroethane (DDE), 152 Dichlorodiphenyltrichloroethane (DDT), 151–152 Dietary-related weight gain, 447–449 Diethylstilbestrol (DES), 26–27 Differential methylation, 442 Dihydrotestosterone (DHT), 41–42, 418 Dioxin, 152–153 Di(2-ethylhexyl) phthalate (DEHP), 151–152, 446 Dll4, 559 DMOT4039A, 559 DMRT1, 298–299 DNA damage, 5–6, 575–576 DNA epigenetic dynamics germ cell development, 211–212, 211f oocyte growth and maturation, 211–213, 211f DNA methylated regions (DMRs), 151 DNA methylation, 151 patterns, 211–212 DNA methyl transferase proteins, 171 Docetaxel, 557, 563 Dominant follicle selection, 55 Donor oocytes and embryos, 503–504 Dosage-sensitive sex reversal/adrenal hypoplasia critical region X (DAX-1), 461 Doxil, 588 Doxorubicin, 563–564 DRE25, 464–465 “Driver” mutations, 548 Drosophila fat facets related X (DFRX), 464 D19S884, 440 Durvalumab, 562

INDEX

E

4E-binding protein 1 (4E-BP1), 32 Echinomycin, 221, 224–225, 258 ECM. See Extracellular matrix (ECM) Edn2, 224–225 Eflornithine, 425 EGF-like factors (Egf-L), 222–223 EGFR. See Epidermal growth factor receptor (EGFR) Egfr expression, 40 Elective single embryo transfer (eSET) cycles, 392 Elimination phase, TME, 583 Embryo banking, 501 Embryo cryopreservation, 503 Embryonic development, 157 Embryonic genome, 161 Embryonic stem cells (ESCs), 504 Embryos in vitro management of, 397 morphological assessments of, 388–390 PGS hypothesis 2.0., 391 blastocyst-stage embryos, 391 clinical utilization, 390 ESTEEM trial, 390–391 ICM lineage, 391 3.0/PGT-A, 391 polar body biopsy, 390 Empty spiracles homeobox 2 (Emx2), 299 EMT. See Epithelial-to-mesenchymal transition (EMT) Enclomiphene, 425–426 Endocrine, 8 Endocrine-disrupting chemicals (EDCs)., 445–447 Endometrial hyperplasia, 421–422 Endometrioid carcinomas (ECs), 515f, 517 Endometrioid ovarian cancer, 536 Endometriomas, 517 Endometriosis, 517 characteristics, 119, 330–331 and TGF-β superfamily members, 119 Endoplasmic reticulum (ER) function, 158–159 redistribution of, 194–195 sterol esters synthesis in, 86 Endothelial cells, 274–275, 582 Endothelin-2 (EDN2), 258 Endothelin-2 and endothelin antagonists block ovulation, 224–225 ENMD-2076, 559 Enoticumab (REGN421), 559 Environmental contaminants, ovarian toxicity of biological plausibility of effects and potential mechanisms, 487–488 biomonitoring studies, 486 ovarian dysfunction, epidemiological evidence of, 486–487 ovarian exposure, 486 EOCs. See Epithelial ovarian cancers (EOCs) EPCAMhi (epithelial) cells, 539–540 EPCAMlo (mesenchymal) cells, 539–540 Epi-A, 536–537 Epi-B, 536–537

Epidermal growth factor (EGF), 40, 306 Epidermal growth factor receptor (EGFR) function, 40 pathway targets, 557–558 Epigenetic marks, 161 Epigenetic maturation epigenetic modifications chromatin immunoprecipitation, 171–172 CpG islands, 171 definition, 170 de novo methylation patterns, 171–172 DNA methyl transferase proteins, 171 genome-wide topological organization, 170–171 germ line differentially methylated region, 171–172 H3K4me3 mark, 172 global silencing, 173 granulosa cells, 170 histone modifications, 173 large-scale chromatin remodeling definition, 173–174 global histone deacetylation, 175 maternal age effect, 175–176 meiotic histone code, 175 NHK-1, depletion of, 174 nonsurrounded nucleolus, 173–174 surrounded nucleolus, 173–174 oocyte meiotic maturation DNA methylation and demethylation, 198–199 epigenetic modifications, GV oocyte, 196–197, 197f histone H3 lysine-4 trimethylation, 198, 199f histone variants, 197–198 polycomb-repressive complexes, 199–200 process of, 170 Epigenetic modifications, 161–162, 170 PCOS definition, 441 differential methylation, 442 hypermethylation, 441 hypomethylation, 442 micro-RNA, 442–443 X-chromosome inactivation, 442 Epigenetic regulation of genome, 161–162 Epigenetic reprogramming, 161 Epigenetics definition, 149 mechanisms, 149 Epigenome, 151, 441–443 Epithelial cell adhesion molecule (Epcam), 516 Epithelialmarkers, 512 Epithelial-mesenchymal plasticity, 530 Epithelial ovarian cancers (EOCs), 512, 514, 521, 547 definition, 514 exosomes, 585 microenvironment., 520 sex cord-stromal tumors adult granulosa cell tumors, 518 sertoli-leydig cell tumors, 518

INDEX

steroid hormone stimulation, 519 subtypes of clear cell carcinoma, 517 endometrioid carcinoma, 515f, 517 high-grade serous carcinomas, 515–516, 515f low-grade serous carcinoma, 515f, 516–517 mucinous carcinoma, 515f, 517–518 type II tumors, 514 type I tumors, 514 VEGF, 522 Epithelial-to-mesenchymal transition (EMT), 521, 529–531, 531f, 536–537, 540 cancer stem cells, 538–539 CD 332, 530 chemoresistance, 538 consequences of, 537–540, 540f epithelial cell-cell junctions, 530 gene expression, 530 immunosuppression, 539–540 inducer, 532 “metastable” states, 530–531 metastasis, 537–538 migration and invasion, 530–531 miR-34 and H19, 530 p120-catenin, 530 and stemness, 531–532 transcription factors, 530 Epoxide hydrolase 1 (EPHX1) gene, 442 Equilibrium phase, TME, 583 ERBB2 mutations, 514, 516–518 Erlotinib, 557 Escape phase, TME, 583 Esr1, 41 ESTEEM trial, 390–391 Estradiol, 9–10, 55–56, 58, 83, 503 Estrogen, 9–10, 41 Estrogen receptor, 487–488 Estrogen receptor-α (Esr1), 26 Estrogen receptor-β (Esr2), 26 Estrogen receptor signaling, 41 Estrogen-related receptor gamma (ERRγ), 487–488 Estrogen-responsive element (ERE), 41 European-ancestry studies, 439 Everolimus, 565 Exogenous androgen, 417 Exosomes, 524, 584–585 Expressed sequence tag (EST), 464–465 Extracellular matrix (ECM), 116, 261–262, 498, 504, 530, 582 Extracellular signal-regulated kinase (ERK) pathway, 556, 556f Extracellular signal-regulated kinases 1 and 2 (Erk1/2), 219–220, 219–220f EZH2, 559–560

F

Factor in the germline alpha (Figla), 303, 303t Fallopian tube epithelium (FTE) origin of ovarian cancer, 529–530, 536 and OSE, 529–530, 532–534, 533t, 537, 541 stemness, 532–534, 533t, 534f Fanconi anemia complementation group L (FANCL), 474

Fanconi anemia family, 303 Farletuzumab, 558 Fatty acid-binding proteins (FABPs), 522 Ferriman-Gallwey scores, 425 Ferrodoxin (FDX1), 237–238 Ferrodoxin reductase (FDXR), 237–238 Fertility preservation after cancer therapy donor oocytes and embryos, 503–504 gestational carrier/adoption, 504 before cancer therapy embryo and oocyte cryopreservation, 503 ovarian tissue cryopreservation and transplantation, 503 contraception options, 504, 505t impact of cancer therapy, minimizing in female fertility, 501, 502t GnRHa, 501 imatinib, 502 sphingosine-1-phosphate (S1P), 502 tamoxifen, 502 menstrual suppression anemia and thrombocytopenia prevention, 504–506 combined oral contraceptives, 506 depot medroxyprogesterone acetate, 506 GnRHa, 504 levonorgestrel intrauterine device, 506 progestin-only pills, 506 ovarian cortical strips, transplantation of, 504 ovarian reserve, 502–503 Fertilization, 157–158 Fetal programming hypothesis, 443 animal-based studies, 443–444 human observational studies, 444–445 mechanisms, 443 Fibroblast growth factor 2 (FGF2), 260–261, 498 Fibrothecomas, 101 FIGLA (POF6), 472 FLIP, 577 Flutamide, 424–425 FMR1 gene, 99–100, 478f Folate receptor-alpha, 558 Follicle(s), 485 activation, 77 atresia, 487, 493 divergence, 55 formation, 76 mammalian, 3–4, 4f ovarian formation and development of, 24f selection of, 4–5 waves, 8 Follicle selection, 5–8 activation of, 6–7 in anovulatory waves, 58–59 death, selection for, 5–6 dynamic changes in, 7–8 intra-and extragonadal hormonal control of, 8–11 gonadotropin-dependent follicle growth and preovulatory follicle selection, 10–11

599 gonadotropin-independent follicle growth, 8–10, 9f oocyte regulation GC lineage differentiation, oocyte paracrine signals regulation, 13–14, 15f GDF9 and BMP15, 11–13, 14f oocyte-secreted factor signaling, impact of, 14–16, 16f oocyte-somatic cell bidirectional communication, 16–17 via key paracrine messengers, 11 ovarian reserve of, 5 Follicle-stimulating hormone (FSH), 4, 8, 23, 53, 60–61, 363, 415, 477, 485, 487, 503 age-specific, 383f β-subunit, 439 mutations, 135–136 polymorphisms, 136 gene structure, 128, 128f three-dimensional structure, 128–130, 128f Follicle-stimulating hormone receptor (FSHR), 38 mutation of activation, 140–141 inactivation, 141–143 location, 137f polymorphisms, 143–144 structure of activation and signal transduction, 132 gene, 131 mRNA, 131 protein, 131–132 TM α-helices organization, 132 Folliculogenesis, 3–4, 107, 113–114, 116, 487 AMH early, 404–405 late, 404–405 production and action, 405f animal models, 62–63 anovulatory waves, follicle selection in, 58–59 antral follicles, recruitment of, 53–54 clinical implications, 63 dominant follicle selection, 55 early stages androgen supplementation, 395–396 HGH supplementation, 396–397 Fanconi anemia family, 303 follicle development and selection, luteal influences, 56–57 follicle divergence, 55 follicle dominance, 56, 57–58f follicular wave patterns, repeatability in, 59 Gas2, 302 GCC breakdown, 300–301 germ cell deletion, 302 gonadotropin sensitive stage description, 392 mild stimulation, 392–393 natural cycle IVF, 391–392, 393f Niche protocols, 394–395, 394t standard stimulation protocols, 392–394, 394t Hes1, 302 Hippo signaling pathway, 302–303

600 Folliculogenesis (Continued) hormonal influences, 60–61, 60f during interovulatory interval, 54–55 language of, 51–52 meiotic arrest, 305–306 mTORC, 301–302 Notch signaling pathway, 302 oocytes, loss of, 300–301 oocyte-specific transcription factors, 303, 303t PDK1-Kit-PI3K/PTEN/AKT-FOXO3 signaling network, 301 PI3K/AKT/mTORC1 signaling, 301, 301f preantral and early antral follicle development, 52–53 preovulatory follicle development, 57–58 prepubertal and pubertal period, 53 primordial follicles formation, 300–301 reproductive senescence, transition to, 59 secondary follicle stage, 303 selection, unified theory of, 55–56 TAF4B, 300–301, 301f TSC1 and TSC2, 301–302 virtual histology approach, 63 Follistatin (FST), 27 abnormalities in PCOS, 101 action of, 97 biosynthesis, 98 FST288, 98–99 FST303, 98 FST315 isoform, 98 gem cells formation, 98–99 inhibits FSH, 95–96 knockout approach, 98 molecular structure, 96 in ovary, 98 structure, 95–96 women with PCOS, 119 Follistatin like-3 (FSTL3), 95–96 Forkhead box L2 (Foxl2), 298 Foxl2 gene, 25, 479, 487 FOXL2 mutation, 518 FOXO3, 29–30 FOXO4, 465 FOXP3, 539 Fragile X (FRAXA), 478–479 Fragile X mental retardation protein (FMRP), 478 Fragile X syndrome, 478 Frasier syndrome, 474 FRAYA. See FMR1 gene "Freeze-all" embryos, 375, 427 Frequently sampled intravenous glucose tolerance test (FSIGT), 420 Frozen-thawed ovarian tissue heterotopic ovarian transplantation, 494–495, 494t, 495–496f, 497 orthotopic ovarian transplantation, 494–497, 494t, 495–496f FSH. See Follicle-stimulating hormone (FSH) FSHR. See Follicle-stimulating hormone receptor (FSHR) FST. See Follistatin (FST) FTY 720, 498 Functional ovarian reserve (FOR), 380

INDEX

G

Galactosemia, 476 Galectins, 278–279 γ-irradiation, 8 γ-secretase inhibitor, 28 Gap junction protein alpha 1 (GJA1), 299 GATA4, 38–39 GATA proteins, 33–34 GCs. See Granulosa cells (GCs) GDF9. See Growth differentiation factor 9 (GDF9) gDMRs. See Germ line differentially methylated regions (gDMRs) Gefitinib, 557 Gelsolin (GSN), 576–577, 578–580f, 582 Gemcitabine, 557–560 Gene expression abnormal expression, 213–214 cholesterol, 92 DNA epigenetic dynamics germ cell development, 211–212, 211f oocyte growth and maturation, 211–213, 211f epithelial-to-mesenchymal transition, 530 during oocyte maturation germinal vesicle, 208 GV-and MII-stage oocytes, 208 meiosis I progresses, 208 miRNA, 210 mRNA stability, 209 oocyte proteome, 210–211 proteasome pathway played, 208 single-cell RNA-seq techniques, 208–209 transcriptome analysis, 208 translation regulation, 210 during oogenesis follicle activation and development, 206–208, 207f molecular mechanisms, 207–208, 207f molecular regulation, 205–206 PGCs’ migration and proliferation, 206 primordial follicle formation, 206 profiles, 512, 530–531 Genome, epigenetic regulation, 161–162 Genome-wide association studies (GWAS), 428–429, 438–440 Chinese-ancestry studies, 439 description, 311 European-ancestry studies, 439 linkage disequilibrium (LD)-based, 311 next-generation sequencing, 322 PCOS across ethnicities, 318 in European populations, 317–318 in Han Chinese population, 311–317 in Korean populations, 317 list, 312–316t POF/I across ethnicity, 319–321 in Dutch population, 319 Han Chinese population, 319 list, 320t SNP-based, 322 Genomic imprints, 161

Germ-cell failure in male (46,XY), 474–476 nest breakdown, and primordial follicle formation, 24–29 growth factors and somatic cell cycle progression, 27–28 neonatal oocyte survival, molecular control of, 25–26 Notch signaling, regulation, 28–29 steroid hormones, 26–27, 26f tumors, 514 Germinal vesicle (GV), 165–166 oocytes, 159–160 stage, 208 Germinal vesicle breakdown (GVB), 165–166 Germ line differentially methylated regions (gDMRs), 171–172 Gestational carrier/adoption, 504 GI50 dose, 538 Global histone deacetylation, 175 Glucose, 420 Glucose regulatory protein 78 (GRP78), 89–90 Glucose transporter 1 (GLUT1), 258 GnRH agonist, 424, 426–427 GOG 213, 554 GOG 218, 553–554 GOG-3005, 553 Gold nanoparticles (AuNPs), 588 Gonadal injury, risk of, 501 Gonadal ridge epithelial-like (GREL) cells, 71–72, 518 Gonadal ridge formation, 72–75, 74f Gonadotoxic therapies, 493–494 Gonadotropin(s), 5, 9, 40, 426, 439, 487, 498. See also Follicle-stimulating hormone (FSH); Luteinizing hormone structure, 127–130, 128f subunits of common α, 132 FSH β, 135–136 genes, 128, 128f, 144 hCG β, 134–135 LHB, 132–134 Gonadotropin-dependent follicle growth, 10–11 Gonadotropin-independent follicle growth, 8–10, 9f Gonadotropin receptor(s) activating mutations, 144 classification, 136 FSHR, 140–141 LHCGR, 138–139 descriptions, 130 inactivating mutations, 144 classification, 136 FSHR, 141–143 LHCGR, 139–140 polymorphisms, 144 FSHR, 143–144 LHCGR, 140 structure, 130–132 Gonadotropin-releasing hormone agonist (GnRHa) endometriosis, 331

601

INDEX

impact of cancer therapy, minimizing, 501 menstrual suppression, 504 Gonadotropin-releasing hormone agonist trigger description, 365 and "freeze-all" embryos, 375 hCG-based triggering, 364–365 LH AND FSH, 365–366, 366f LPS post with hCG, 369–372, 370–373f intense, 369, 370f with LH, 373, 374f mid-luteal phase, 369 nasal buserelin three times a day, 373–374, 374f to ovarian responses, 374 luteolysis, 367–368 mid-cycle ovulation, 363–364 oocytes retrieved and embryo quality, 366–367 preparation, 366 Gonadotropin-releasing hormone antagonists in luteal phase, 354 ovarian stimulation protocols (see Gonadotropin-releasing hormone agonist trigger) for ovulation suppression, 351 protocols, 351, 354 Gonadotropin stimulation, 392 G-protein-coupled receptor (GPCR), 38–39 G protein-coupled receptor 30 (Gpr30), 41 Graafian follicles, 5, 487 Granulosa cells (GCs) acquisition of, 34–35 AKT/PI3K signaling pathway, 40 bovine ovarian development, 71–72 cumulus, 165 epigenetic maturation, 170 of ER-β-knockout follicles, 41 nuclear maturation mammalian oocyte maturation, 165 mammalian oogenesis, 159–160 origin of, 71–72 proliferation, 38–42 FSHR signaling and kinase cascades, 39–40 regulation of, 36–37 steroid hormones, in follicular maturation, 40–42 structure and function, 3–4, 4f transcription and chromatin remodeling, 170 Green fluorescent protein (Gfp), 28 Gremlin-1 (Grem1), 298 Growing follicles, 5–7 Growth arrest-specific gene 2 (Gas2), 302 Growth differentiation factor 9 (GDF9), 11–13, 14f, 35–36, 160, 487 Growth differentiation factors (GDFs), 107 Growth factors, 5, 11, 27–28 Growth-regulated oncogene-a (GROa), 522 Gut-derived endotoxinemia, 449 Gut microbiota, 449–450

GV. See Germinal vesicle (GV) GWAS. See Genome-wide association studies (GWAS) Gynecologic cancer, 421–422

H

Hairy and enhancer of split 1 (Hes1), 302 Hanna (FecXH) mutation, 35–36 hCG β-subunit (CGB) mutation, 134–135 polymorphisms, 134–135 structure, 128 HDLs. See High-density lipoproteins (HDLs) Hematological disorders, 493 Hematopoietic stem cell and replacement therapies, 338 Hematopoietic stem cell transplantation (HSCT), 498 Heme oxygenase 1, 258 Hemoglobin A1c (HbA1c), 420–421 HER2, 557–558 Hes1, 37–38 Heterodimerization, 557 Heterotopic ovarian transplantation, 494–495, 494t, 495–496f, 497 Hexokinase II (HKII), 577–578 HFM1 (POF9), 472 HGSC. See High grade serous carcinoma (HGSC) Hierarchical stem cell model, 532 HIF1. See Hypoxia-inducible factor 1 (HIF1) High-density lipoprotein receptors, 85–86 High-density lipoproteins (HDLs), 85 Higher risk disease, 553–554 High-grade endometrioid, 514 High grade serous carcinoma (HGSC), 514–516, 515f, 518–520, 536–537, 547–548 Hippo signaling pathway, 302–303 Hirsutenone, 587–588 Hirsutism, 418–419, 421–425 Histone acetyltransferases and deacetylates (HDAC), 560 Histone deacetylase inhibitor (HDACi) treatment, 587 HMGCR, 247–248 Homodimerization, 557 Homologous recombination (HR), 549 Homologous recombination defective (HRD), 551–552 Hormone-sensitive lipase (HSL), 241–242 HSD3B1/2, 246 HSD17B4, 473 Human chorionic gonadotropin (hCG), 128–130, 494 Human Genome Project, 427 Human inhibitor of apoptosis protein 1 (HIAP1 or cIAP-2), 578–579 Human inhibitor of apoptosis protein 2 (HIAP2 or cIAP-1), 578–579 Human mtDNA, 327 Hyaluronan synthesis, 117 Hydrocarbon (JP-8), 153 5-Hydroxytestosterone (5-HT), 448 Hyperandrogenemia, 418–420, 424

Hyperandrogenism, 41–42, 417–418, 422, 440–441, 448 Hypergonadotropic hypogonadism, 473 Hyperinsulinemia, 415, 418–420, 424, 427, 448 Hypermethylation, 441 Hyperprolactinemia, 415–416 Hyperreactioluteinalis, 348 Hyperthecosis, 417 Hypogonadotropic hypogonadism, 100, 502 Hypomethylation, 442 Hypothalamic-pituitary-ovarian (HPO) axis, 415 Hypothalamopituitary adrenal (HPA) axis, 448 Hypothyroidism, 415 Hypoxia, 332, 332f Hypoxia-inducible factor 1 (HIF1) HIF1 activation, 256–257 HIF1A-dependent genes, 256–258, 256–257f independent regulation, 258–259 LH-dependent gene expression, 256, 257f mechanisms, 256, 256f microRNA-210, 259

I

ICON7, 553–554 IL6-STAT3-HIF pathway, 548 Imatinib, 502, 561 Immature oocyte cryopreservation, 501 Immune cell infiltration and inflammatory mediators leukocyte infiltration, 225 NO and ROS, 225–226 prostaglandins, 226 Immune cells CCL2, 277 CD5+ and CD8+ cells, 276 eosinophils, 277 gene expression arrays, 276 luteal resident T cell function, 276 macrophage infiltration, 276–277 neutrophils, 277 WC1+ gamma delta T cells, 276 Immune checkpoint blockers, 586–587 Immunosuppression, 539–540 Immunotherapy, 561–562, 562f, 584 Impaired glucose tolerance (IGT), 420 Inactivating luteinizing hormone receptor defect (46,XX), 481 Infertility, 421, 486–487 risk factors, 485 treatment of clomiphene citrate, 425–426 gonadotropins, 426 in vitro fertilization, 426–427 laparoscopic ovarian drilling, 426 letrozole, 426 lifestyle modification, 425 Inflammation, 535–536 Inflammatory mediators leukocyte infiltration, 225 NO and ROS, 225–226 prostaglandins, 226 Inhibin, 36 A, 95 pregnancy and labor, 100

602 Inhibin (Continued) regulation during human menstrual cycle, 97 α-subunit, 96–97 B, 95, 487, 503 activity in puberty, 100 autoimmune oophoritis, 100, 102 decreased follicular-phase FSH levels, 99 follicle development, 101 ovarian aging, 99 Prader Willi syndrome, 100 in primary ovarian insufficiency, 99–100 regulation during human menstrual cycle, 97 STRAW staging system, 99 β promoter, 97 estradiol vs., 97 FSH biosynthesis, 97–98 human physiology and clinical implications FSH regulation for controlled follicle development, 99 in vitro fertilization, 100–101 menopause, 99 ovarian aging, 99 ovarian cancers, 101 ovarian follicle reserve, 100–101 PCOS, 101 perimenopause, 99 POI, 99–100 pregnancy and labor, 100 in puberty, 100 molecular structure, 96 purification of, 95–96 receptor, 96 sequence for, 95 signaling, 96 structure, 95 terminology, 95 Inhibitor of apoptosis proteins (IAP), 576 and chemoresistance, 578–581 X-linked inhibitor of apoptosis protein, 578–582 Inner cell mass (ICM) lineage, 391 Insulin growth factor (IGF), 61 Insulin growth factor receptor (IGFR), 40 Insulin inhibits aromatase activity, 445 Insulin-like growth factor-1 (Igf1), 304 Insulin-like growth factors, 487, 535 Insulin resistance, 419–420, 448–449 Intact GSN (I-GSN) protein content, 579f Interferon-stimulated genes (ISGs), 278–279 Interleukin 6 (IL-6), 520–521 International Cancer Genomics Consortium, 547–548 Interovulatory interval (IOI), 52, 54–55 Intracellular signaling cascades cAMP-dependent serine protein kinase A (PKA) activation, 218–219 CCAAT/enhancer binding proteins alpha and beta, 219–221 Gαq/11/PKC pathway, 219 in granulosa cells, 219 intracellular signaling pathways, 218–219, 219f LH surge, 217–218, 218f

INDEX

Intrauterine nutritional insufficiency, 445 Inverdale (FecXI) mutation, 35–36 In vitro fertilization (IVF), 426–427, 497, 504 ART adverse outcomes, 383–384 age dependent, 384 multiple pregnancies, 383–384 offering centers, 384 for oldest patient population, 384–385 U.S. centers report, 384 U.S. live birth rate, 384, 384f description, 363 failure by autologous mitochondrial injection, 329 inhibin B, 100–101 Kp-54 administration, 352–353 natural cycle, 391–392, 393f outcomes clinical pregnancy and live birth rates, 388, 389f embryos quality assessments, 388–391 quantity and quality of oocytes, 388 In vitro fertilization–intracytoplasmic sperm injection (IVF–ICSI), 339 In vitro follicle maturation, 504 Ipilimumab, 561, 586 I-SPY2 breast cancer trial, 552 IVF. See In vitro fertilization (IVF)

J

Jag1, 28–29 Jagged1 ligand (Jag1), 302 Janus kinase, 520 JP-8 (hydrocarbon), 153

K

Kisspeptin, 385, 426–427 Kit ligand (KITL), 6–7, 27 Knockin mouse genetic models, 296 Knockout (KO) mouse genetic models, 98, 118, 296 KRAS mutations, 514, 516–518, 548, 556, 565 Kynurenine, 583

L

Lactobacillus, 449–450 Lapatinib, 557–558 Laser microdissection, 205–206 LDLR. See Low-density lipoprotein receptor (LDLR) Leaky gut hypothesis, 449 Leptin, 521–522 Letrozole, 426, 449, 565 Leukocyte infiltration, 225 Levonorgestrel intrauterine device (IUD), 506 Lgr5, 514–515, 535 LGSCs. See Low-grade serous carcinomas (LGSCs) LH. See Luteinizing hormone (LH) LHCGR. See Luteinizing hormone/chorionic gonadotropin receptor (LHCGR) LH-mediated cell cycle resumption, 160 LHR. See Luteinizing hormone receptor (LHR) LHX8, 33

Lhx8 (LIM homeobox8), 303, 303t LIM class homeodomain protein, 512 Lim homeobox genes 1 and 9 (Lim1 and Lhx9), 299 Lim-homeobox protein, 33 LIN28A, 33 Lipases hormone-sensitive lipase or lipase E (LIPE), 237–238, 248. See also Hormonesensitive lipase (HSL) Liposomal doxorubicin, 557 Liposomal lurotecan, 588 Liposomal topotecan, 588 Livin, 578–579 Long Evans Hooded vs. Sprague-Dawley rats, 488 Loss of heterozygosity (LOH), 551 Low-density lipoprotein receptor (LDLR), 85–88, 247 Low-density lipoproteins (LDLs) pathway, 84–85 Low-dose estradiol with progestin, 504 Low functional ovarian reserve (LFOR), 380 Low-grade endometrioid carcinoma, 514, 517 Low-grade serous carcinomas (LGSCs), 514, 515f, 516–517 Low-grade serous ovarian cancers, 548 Low malignant potential (LMP serous tumors), 516–517, 560 Lunatic fringe (Lfng), 28 Luteal angiogenesis bovine ovary 1-day postovulation, 260, 260f FGF2, 260–261 PGE2, 261 VEGFA, 260–261 Luteal phase dominant follicles (LPDFs), 59 Luteal-phase support (LPS), 364–365. See also Gonadotropin-releasing hormone agonist trigger Luteal regression, 238 Luteal rescue during early pregnancy, 284 IFNT, 285 maternal recognition period, 285 PAG, 285 pig, 283 primate, 281–282, 283f rodent, 282–283, 283f in ruminant IFNT, antiluteolytic actions of, 279–280 IFNT, direct actions of, 280–281 mechanisms, 283f Luteinizing hormone (LH), 4, 23, 53, 61, 160, 160f, 363, 415, 439, 485, 487, 534–535 Luteinizing hormone/chorionic gonadotropin receptor (LHCGR), 238 function, 127 mutation of activation, 138–139 inactivation, 139–140 location, 137f polymorphisms, 140 structure of activation and signal transduction, 131 gene, 130 messenger ribonucleic acid, 130 protein, 130–131

INDEX

three-dimensional, 128f, 131 TM α-helices organization, 131 Luteinizing hormone receptor (LHR) β-subunit mutations, 132–133 polymorphisms, 133–134 structure, 128 on cumulus cells, 160f expression, 8–9 knockout mice, 8–9, 133, 140 mutations in 46,XX, 481 Luteogenesis, 61–62 Luteolysis cell loss, 279 initiation of, 271–272 luteolytic actions cytokines, 277–278 endothelial cells, 274–275 immune cells, 276–277 steroidogenic cells, 272–274 luteolytic capacity, acquisition of, 270–271, 272f molecular mediators, 278–279 Luteolytic actions cytokines, 277–278 CXCL8, 278 FAS-FAS ligand system, 277–278 IFNG, 277–278 matrix remodeling, 278 production and regulation, 277 TNF, 277 transcription, 277 endothelial cells, 274–275 immune cells CCL2, 277 CD5+ and CD8+ cells, 276 eosinophils, 277 gene expression arrays, 276 luteal resident T cell function, 276 macrophage infiltration, 276–277 neutrophils, 277 WC1+ gamma delta T cells, 276 steroidogenic cells ATF3, 273 downstream effects, 273, 274f induced progesterone production, 272–274 in vitro studies, 272 mechanism of action, 273 Luteolytic capacity, acquisition of, 270–271, 272f Luteotrophin, 273–274 LY90009, 559 Lysyl oxidase (LOX), 116

M

Macrophage infiltration, 276–277 ovulation, 225 Major histocompatibility complex (MHC), 583–584 Major histocompatibility complex class I (MHCI) receptors, 562f Mammalian follicles, 3–4, 4f Mammalian oocyte, 3

Mammalian oocyte maturation epigenetics (see Epigenetic maturation) nuclear maturation cell cycle molecules, 166–168, 167f germinal vesicle, 165–166 granulosa cells, 165 LH, induction of, 170 meiosis, 165 meiotic competence, 168 meiotic resumption, 165–166 mouse oocytes, 165–166, 166f preantral-antral stage, 165 primordial germ cells, 165 prophase-I arrest, 165–166, 168–169 Mammalian oogenesis cytoplasmic maturation early embryonic development, 157–158 fertilization, 157–158 maternal effect genes, 158 reorganization, 158–159 epigenetic regulation, 161–162 nuclear maturation bone morphogenic factor 15, 160 CDC25, active/inactive, 160 cell cycle, 159–160, 159f germinal vesicle oocytes, 159–160 granulosa cells, 159–160 growth differentiation factor 9, 160 mechanism, 159–160 meiotic resumption, 160, 160f metaphase arrest, 160–161 primordial follicles, 159–160 prophase arrest CDK1 protein, 160 primordial germ cells, 157 Mammalian ovary, 3 development of (see Ovarian development) estrogen receptor, 41 follicle selection in (see Follicle selection) organization, 296 primordial follicles, 208 Mammalian target of rapamycin (mTOR), 32 Mammalian target of rapamycin complex (mTORC), 301–302 Mammalian target of rapamycin complex 1 (mTORC1), 207 MAP kinase phosphatase 3 (MKP3), 40 MAPKs. See Mitogen-activated protein kinases (MAPKs) Massive parallel sequencing, 213–214 Mater. See NOD-like receptor family pyrin domain containing 5 (Nlrp5) Maternal effect genes, 158 Maternal mitochondrial genome (mDNA), 379 Maternal recognition of pregnancy (MRP), 279 Matrix metalloproteinase 11 (MMP11), 521 Matrix metalloproteinases (MMPs), 262 Matrix remodeling, PGE2, 261–262 Maturation promoting factor (MPF), 159f, 166–167, 182, 305–306 MCM8 (POF10), 472 MCs. See Mucinous carcinomas (MCs) MD (myotonic dystrophy), 479 MDSCs (myeloid-derived suppressor cells), 583

603 Mechanical hair removal, hirsutism treatment, 425 Mechlorethamine, 503 Meiosis definition, 165 initiation, 165 process of, 157 Meiotic arrest, 305–306, 305f Meiotic cell cycle, 157, 159–160 Meiotic resumption, 160, 160f, 165–166 MEK inhibitor, 521, 556, 565 Melatonin, 498 Menopause age at, 480 early, 476, 502 inhibin B, 99 reverse, 498–499 STRAW staging system, 99 Menstrual cycle, 487, 493 disruption of, 486 function, 486 luteal phase of, 485 ovarian reserve, 502 Menstrual suppression anemia and thrombocytopenia prevention, 504–506 combined oral contraceptives, 506 depot medroxyprogesterone acetate, 506 GnRHa, 504 levonorgestrel intrauterine device, 506 progestin-only pills, 506 Mes, 536–537 Mesenchymal components, 512 Mesenchymal phenotype, 530 Mesenchymal stem cells (MSC), 520–521 Mesenchymal-to-epithelial transition (MET), 531–532, 533t Mesoderm, 157 Mesothelin, 559 Mesothelium, 512 MET. See Mesenchymal-to-epithelial transition (MET) Metaanalysis gene-set enrichment of variant associations (MAGENTA) analysis, 322 “Metastable” states, 530–531 Metformin, 351–352, 424 Methoxychlor, 151–152, 487 Methylglyoxal (MG), 453 Metronomic cyclophosphamide, 563 MGCs. See Mural granulosa cells (MGCs) Microparticles (MPs), 585 MicroRNA-210, 259 MicroRNA (miRNA), 441–443, 518, 523–524, 537 Microsatellite instability (MSI), 561 miR-6126, 524 miRNA. See MicroRNA (miRNA) miRNA-200 family, 537, 539 Mitochondria, 158 abnormal function, 341 autologous germline transfer, 340 coenzyme Q10 supplementation, 339 DNA at blastocyst stage, 340–341 at cleavage stage, 340

604 Mitochondria (Continued) oocytes and CC, 340 in endometriosis, GnRH agonist treatment, 330–331 low oxygen tension and embryo development blastocysts cultured under, 331–332, 332f hypothetical model, 332, 332f MtMP, 331–332 in oocytes, 338 ooplasmic transfer in mature human oocytes, 339–340 origin, 337 replacement therapy, 338 reproductive aging, 339 role, 329, 333 segregation, 337–338 Mitochondrial deoxyribonucleic acid (mtDNA) in assisted reproduction cytoplasmic transfer, 327–329 mitochondrial disorders, 327 preimplantation genetic diagnosis, 327 GnRH treatment on endometriosis, 330–331 human, 327 and human embryo viability copy number, 329 double embryo transfer, 330 euploid blastocysts, 330 in euploid embryos, 329–330 quantity, 329 mutations, 327 Mitochondrial disorders, 327 Mitochondrial membrane potential (MtMP), 331–332 Mitochondrial RNA, 329 Mitogen-activated protein (MAPK)-ERK kinase (MEK) pathway, 556, 556f Mitogen-activated protein kinases (MAPKs), 39–40, 39f, 166–167, 167f, 185 Mitogen-activated protein kinases 3 and 1 (MAPK3/1). See Extracellular signalregulated kinases 1 and 2 (Erk1/2) Mitosis, 157 Mixed epithelial/mesothelial phenotype, 512 MMP9, mRNA-expression of, 262 Modified Ferriman-Gallwey score, 419 MOFs (multi-oocytic follicles), 26–27 Monosomy X, 462 Mouse oocytes, 165–166, 166f mRNA stability localization, 209 poly(A) tail length and 3 terminal uridylation, 209 ribonucleoprotein particles, 209 subcortical maternal complex, 209 MSH5 (POF13), 473 mtDNA. See Mitochondrial deoxyribonucleic acid (mtDNA) MtMP. See Mitochondrial membrane potential (MtMP) mTOR pathway, 555–556, 555f Mucinous carcinomas (MCs), 514–515, 515f, 517–518

INDEX

M€ ullerian ducts, 512, 513f, 514–516, 518, 529–530 Mullerian-inhibiting substance (MIS). See AntiM€ ullerian hormone (AMH) Multi-oocytic follicles (MOFs), 26–27 Mural granulosa cells (MGCs), 13, 15, 165 Myeloid-derived suppressor cells (MDSCs), 583 Myotonic dystrophy (MD), 479

N

Nab-paclitaxel, 563 N-acetylcystein, 498 Nanog and octamer-binding transcription factor 4 (OCT4), 299 Nanomedicine, 588 NANOS3, 474–475 Nanotechnology, 588 Natriuretic peptide precursor type C (NPPC), 16–17, 169 Natriuretic peptide receptor 2 (NPR2), 169 Natural killer cells (NK cells), 583–584 Neiman-Pick Type C disease, 84–85 Neo-adjuvant chemotherapy, 584 Neovasculature and angiogenesis, 522–524 Neuronal apoptosis inhibitory protein (NAIP), 578–579 Neurotrophic tropomyosin-related kinase (Ntrk) receptors, 27 Neutral cholesterol ester hydrolase 1 (NCEH1), 237–238, 248 Neutrophils, 225 Next-generation sequencing (NGS), 329, 428–429 Nicotine, 447 Nintedanib, 564 Niraparib, 552 Nitric oxide (NO), 225–226 Nitric oxide synthase (NOS), 258, 306 Nivolumab, 561, 586 Nobox (newborn ovary homeobox), 33, 303, 303t NOBOX (POF5), 466–471 NOBOX gene, 118–119 NOD-like receptor family pyrin domain containing 5 (Nlrp5), 158 Noncanonical imprinting mechanism, 173 Nonclassical congenital adrenal hyperplasia (NCCAH), 416 Noncoding RNA, 151 Nongrowing follicles, 3–4 Nongynecological metastases, 519 Nonsurrounded nucleolus (NSN), 173–174 Norethindrone acetate, 504 Notch pathway, 559 Notch receptor intercellular domain (NICD), 28 Notch signaling, 28–29 cellular signals, during preantral follicular development, 37–38 pathway, 206–207, 302 regulation, 28–29 NOVA studies, 551–552 NR5A1 (POF7), 472 N-terminus cytoplasmic gelsolin (N-cGSN), 576–577

Nuclear genome transfer, 328 Nuclear haploid genomes (nDNA), 379 Nuclear maturation bone morphogenic factor 15, 160 CDC25, active/inactive, 160 cell cycle, 159–160, 159f germinal vesicle oocytes, 159–160 granulosa cells, 159–160 growth differentiation factor 9, 160 mammalian oocyte maturation cell cycle molecules, 166–168, 167f germinal vesicle, 165–166 granulosa cells, 165 LH, induction of, 170 meiosis, 165 meiotic competence, 168 meiotic resumption, 165–166 mouse oocytes, 165–166, 166f preantral-antral stage, 165 primordial germ cells, 165 prophase-I arrest, 165–166, 168–169 mechanism, 159–160 meiotic resumption, 160, 160f metaphase arrest, 160–161 oocyte meiotic maturation cyclin B1 levels, regulation of, 184 first meiotic division, 185f, 186–187 high cAMP level, maintenance of, 182–184, 183f homologous chromosome separation, 187, 188f interkinesis, 187–188 low MPF activity, maintenance of, 183f, 184 meiotic resumption, 185–186, 186f MII arrest, maintenance of, 188–189, 189f prophase I arrest, 181–182, 182f protein phosphatases, 184–185, 185f primordial follicles, 159–160 prophase arrest CDK1 protein, 160 Nuclear receptor subfamily 5 group a member 1 (Nr5a1), 299 Nucleolar protein nucleoplasmin 2 (Npm2), 174 Nucleosomal histone kinase-1 (NHK-1), 174

O

Obesity, 421–423, 427, 437, 440, 448 Occult primary ovarian insufficiency (oPOI), 381 OHSS. See Ovarian hyperstimulation syndrome (OHSS) Olaparib, 551, 558–559, 562 Olaratumab (IMC-EGE), 560–561 Omental adipose-derived MSCs (O-ADSCs), 521 Omentum, 520 Oncofertility contraception options, 504, 505t fertility preservation in cancer setting donor oocytes and embryos, 503–504 embryo and oocyte cryopreservation, 503 gestational carrier/adoption, 504 ovarian cortical strips, transplantation of, 504

INDEX

ovarian tissue cryopreservation and transplantation, 503 impact of cancer therapy, minimizing in female fertility, 501, 502t GnRHa, 501 imatinib, 502 sphingosine-1-phosphate, 502 tamoxifen, 502 menstrual suppression anemia and thrombocytopenia prevention, 504–506 combined oral contraceptives, 506 depot medroxyprogesterone acetate, 506 GnRHa, 504 levonorgestrel intrauterine device, 506 progestin-only pills, 506 ovarian reserve, 502–503 “One size fits all” approach, 547, 588 Online Mendelian inheritance in man (OMIM), 465–466 Oocyte aneuploidy, 487 banking, 501 cryopreservation, 503 derived factors, 487 in vitro management of, 397 maturation of inhibitor, 16–17 in vitro fertilization, 363 in vivo fertilization, 363 mid-cycle luteinizing hormone, 363–364 mitochondria in, 338 quantity and quality, 388 scoring system, 390f timing of retrieval, 397 Oocyte meiotic maturation cytoplasmic molecular maturation BTG4 and CCR4-NOT RNA deadenylase, 190–192, 191f dormant mRNAs, translational activation of, 189–190 maternal mRNA decay, 190 ZAR1 and ZAR2, 192 zinc finger protein 36 like 2, 192 cytoplasmic organelle maturation cortical granule migration, 193–194 cytoplasmic maturation, improvement of, 195–196 cytoplasmic quality, evaluation of, 195–196 cytoskeleton dynamics, 195 ER and golgi complex, redistribution of, 194–195 mitochondrial number and distribution, 192–193, 194f epigenetic maturation DNA methylation and demethylation, 198–199 epigenetic modifications, GV oocyte, 196–197, 197f histone H3 lysine-4 trimethylation, 198, 199f histone variants, 197–198 polycomb-repressive complexes, 199–200

nuclear maturation cyclin B1 levels, regulation of, 184 first meiotic division, 185f, 186–187 high cAMP level, maintenance of, 182–184, 183f homologous chromosome separation, 187, 188f interkinesis, 187–188 low MPF activity, maintenance of, 183f, 184 meiotic resumption, 185–186, 186f MII arrest, maintenance of, 188–189, 189f prophase I arrest, 181–182, 182f protein phosphatases, 184–185, 185f Oocyte-secreted factor (OSF) signaling, 14–15 Oocyte-specific constitutive activation of PI3K (PI3KCA), 30–31 Oocyte-specific transcription factors, 303, 303t Oogenesis, 485 Oogonial stem cells (OSCs), 329 OR. See Ovarian reserve (OR) Oral glucose tolerance testing (OGTT), 420–421 Oregovomab (antiCA125), 561 Ornithine decarboxylase, 425 Orthotopic ovarian transplantation, 494–498, 494t, 495–496f OSE. See Ovarian surface epithelium (OSE) Ostradiol, 449 Ovarian autotransplantation techniques cancer cell reimplantation, risk of, 498 frozen-thawed ovarian tissue heterotopic ovarian transplantation, 494–495, 494t, 495–496f, 497 orthotopic ovarian transplantation, 494–497, 494t, 495–496f limitations with, 497–498 ovarian tissue crypreservation, 493–494, 494t success rate of, 497 Ovarian cancer (OVCA) angiogenesis, 553–555 antimesothelin antibodies, 559 Aurora kinases, 559 cell plasticity, 529 chemoresistance, 575–576, 577f cancer-derived extracellular vesicles, 585 cell-based immunotherapies, 587 cellular mechanisms, 576 and cisplatin action, mechanism of, 575–576 cross-talk molecules in TME, 584–586 cytokines/chemokines, 586 gelsolin (GSN), 576–577, 578–580f, 582 hexokinase II, 577–578, 582 immune checkpoint blockers, 586–587 immunotherapy, 584 inhibitor of apoptosis proteins, 578–581 nanotechnology/nanomedicine, 588 natural food compounds, 587–588 P53, 577–578, 582 personalized cancer therapy, 588 PI3-K/Akt pathway, 581–582 plasma gelsolin, 585 targeting epigenetic changes, 587 TILs, chemotherapy on, 584

605 tumor-derived extracellular vesicles, 584–585 tumor microenvironment, 576–584 combination therapy, 562–565 and emerging antiangiogenic therapies, 563–565 PI3K/AKT and MEK inhibitor, 565 cyclin-dependent kinases, 560 EGFR pathway, 557–558 epithelial-to-mesenchymal transition, 530–531, 531f, 540 cancer stem cells, 538–539 CD 332, 530 chemoresistance, 538 consequences of, 537–540, 540f epithelial cell-cell junctions, 530 gene expression, 530 immunosuppression, 539–540 mesenchymal gene expression, 530 “metastable” states, 530–531 metastasis, 537–538 migration and invasion, 530–531 miR-34 and H19, 530 p120-catenin, 530 and stemness, 531–532 transcription factors, 530 folate receptor-alpha, 558 heterogeneity of, 536–537 histone acetyltransferases and deacetylates, 560 homologous recombination, 549–553 immunotherapy, 561–562, 562f incidence rate, 511 inhibins and, 101 lethality of, 575 microenvironment, 519 angiogenesis and neovasculature, 522–524 cancer-associated fibroblasts, 520 in disease progression, 520–525, 525f mesenchymal stem cells, 520–521 ovary-associated adipocytes, 521–522 molecular aberrations in, 547–549, 548t M€ ullerian ducts, 512, 513f Notch pathway, 559 ovarian epithelial cancers clear cell carcinoma, 517 definition, 514 endometrioid carcinoma, 515f, 517 high-grade serous carcinomas, 515–516, 515f low-grade serous carcinoma, 515f, 516–517 mucinous carcinoma, 515f, 517–518 sex cord-stromal tumors, 518 type II tumors, 514 type I tumors, 514 ovarian surface epithelium, 512 ovary, development of, 512 p53, 558–559 phenotypic plasticity ovarian surface epithelium, 529–530 in ovulatory wound repair, 534–536 PI3K/AKT/mTOR pathway, 555–556, 555f platelet-derived growth factor, 560–561

606 Ovarian cancer (OVCA) (Continued) poly-ADP-ribose polymerase inhibitors BRCA-mutated tumors, 549 categories, 549 clinical trials, 549, 550t and DNA damage repair, 549, 550f niraparib, 552 olaparib, 551 rucaparib, 551–552 side effects, 549 talazoparib, 553 veliparib, 552–553 RAS/RAF/MEK/ERK pathway, 556, 556f side effects for targeted therapies, 565 SRC-family kinases, 560 stemness in fallopian tube epithelium, 532–534, 533t, 534f in ovarian surface epithelium, 532, 533t subtypes and origins, 514–518, 514f SWI/SNF chromatinremodeling complex, 559–560 Ovarian cancer-associated EVs, 522–524 Ovarian cortical strips, transplantation of, 504 Ovarian cortical tissue, 495–498 Ovarian development basement membrane, 77 bovine basal lamina, 75 granulosa cells, 71–72 follicle activation, 77 gonadal/genital ridge formation, 72–75, 73–74t ovigerous cords breakdown and follicle formation, 76 formation, 75–76 primordial follicle reserve, 76–77 regionalizaiton into cortex and medulla, 76 schematic diagram, 74f surface epithelium formation, 77–78 thecal interna and externa formation, 78 tunica albuginea formation, 78 vascularization, 78–79 Ovarian dysfunctions. See Polycystic ovary syndrome (PCOS); Premature ovarian failure/insufficiency (POF/I) Ovarian factor aging concept of, 380 physiology of, 381–383 in ART definition, 380–381 IVF treatment (see In vitro fertilization (IVF)) controlled by endocrine and immune systems, 387 GnRH, 385 hypothalamic-pituitary-adrenal axis, 386–387 definition, 379–381 Ovarian failure mendelian causes aromatase mutations (CYP 19), 477 autosomal-dominant POF, 479

INDEX

blepharophimosis-ptosis-epicanthus (FOXL2), 479 carbohydrate-deficient glycoprotein, 476 deficiency of 17a-hydroxylase/17,20desmolase deficiency (CYP17), 476–477, 477f female (46,XX) sibs, 474–476 fragile X syndrome, 478 galactosemia, 476 germ-cell failure in male (46,XY), 474–476 myotonic dystrophy, 479 pleiotropic mendelian disorders, 474 triplet nucleotide repeats, 478, 478f triplet repeats, 479 XX gonadal dysgenesis, 465–473, 465t 46,XY agonadia, 477–478 ovarian development, embryology of, 461 polygenic factors in POF, 480 X chromosome, haploinsufficiency for monosomy X, 462 nomenclature, 462, 462t X long-arm deletions, 464 Xp gonadal determinants, candidate genes for, 464 Xq gonadal determinants, candidate genes for, 464–465 X short-arm deletions, 462–463, 463f Ovarian follicles formation and development of, 24f selection of, 4–5 Ovarian folliculogenesis, 440–441 Ovarian function androgens role in, 10 controlled by endocrine and immune systems, 387 GnRH, 385 hypothalamic-pituitary-adrenal axis, 386–387 disorders, GWAS of (see Genome-wide association studies (GWAS)) mutations and polymorphisms in (see Gonadotropin(s); Gonadotropin receptor(s)) regulation of, 107, 109, 114f, 118 transgenic mouse models (see Transgenic mouse models) ultrasound study, 54 Ovarian hyperstimulation syndrome (OHSS), 363, 382–383, 407, 418 after egg retrieval albumin, 353–354 calcium, 354 combination of treatments, 354 dopamine agonists, 353 GnRH analogs in luteal phase, 354 letrozole, 354 oocyte/embryo cryopreservation, 353 risk factors, 348 causes, 345 clinical manifestation ascites, 349 classification, 349, 349t early and late onset after hCG administration, 349

processes, 348–349 thromboembolic complications, 350 during COS coasting, 352 kisspeptin (Kp), 352–353 lowering/withholding hCG, 352 ovulation triggering with GnRH agonist, 352 risk factors, 348 before COS begins adjuvant therapies, 351–352 recommended stimulation protocol, 351 risk factors, 348 definition, 345 diagnosis, 350–351 economic burden, 345 GnRHa triggering for preventing, 367 incidence, 345 management, 354–357, 355f mild, 356 moderate, 356 severe and critical, 356–357 pathogenesis dopamine receptors expression, 346–348, 347f estradiol levels, 346 granulosa-luteal cells expression, 346, 347f spontaneous OHSS, 348 trophoblast-derived hCG, 345–346 vascular endothelial growth factor, 346 women with PCOS, 346, 347f risk factors, 348t after ovum pickup, 348 during COS, 348 before COS begins, 348 signs and symptoms, 345, 350 Ovarian reserve (OR), 3 age-specific FSH and AMH levels, 382, 383f definition, 502 embryonic development, 157 functional, 380 growing follicle pool, 380 low functional, 380–381 markers anti-M€ ullerian hormone, 503 antral follicle counts, 503 estradiol, 503 follicle-stimulating hormone, 503 inhibin B, 503 menstrual cycle, 502 ovarian volume, 503 normal vs. abnormal, 381–382 ontogeny of, 382f ovarian aging, physiology of, 381–383 ovarian stimulation, 382–383 resting pool of primordial follicles, 380 Ovarian resistance inactivating luteinizing hormone receptor defect (46,XX), 481 XX gonadal dysgenesis, FSHR gene mutation (C566T), 480–481 Ovarian stimulation, 493 Ovarian stroma, 512, 515–516, 519

INDEX

Ovarian suppressive therapies, 424 Ovarian surface epithelium (OSE), 77–78, 512, 513f, 516, 518, 529–530 luteinizing hormone, 534–535 phenotypic plasticity, 529–530 reepithelialization, 535 stemness identification and characterization, 532 regulation of, 532, 533t Ovarian tissue cryopreservation, 501, 504 ovarian autotransplantation techniques, 493, 497 cancer and noncancer indications for, 494t fertility preservation by, 498 history of, 493–494, 494t transplantation, oncofertility, 503 Ovarian toxicity of environmental contaminants biological plausibility of effects and potential mechanisms, 487–488 biomonitoring studies and ovarian exposure, 486 ovarian dysfunction, epidemiological evidence of, 486–487 Ovarian volume, 503 Ovary activating mutation of FSHR, 141f development of, 512 formation, 72f role in mammals, 71 Ovary-associated adipocytes, 521–522 OVCA. See Ovarian cancer (OVCA) Oviductspecific glycoprotein1, 516 OVOL, 531–532 Ovulation, 61–62, 534–535 clinical aspects anovulation, 226–227 contraception, 227–228 COC expansion, 217–218 activation, 220f, 221–222 extracellular matrix production, 222 granulosa cell EGF-like factors, 222–223 oocyte meiosis, resumption of, 221–222 ovulatory pathways, 223 cumulus-oocyte complex, 306 cyclooxygenase pathway, 306 EGF network, 306 extracellular signal-regulated kinases 1 and 2, 219–220, 219–220f immune cell infiltration and inflammatory mediators leukocyte infiltration, 225 NO and ROS, 225–226 prostaglandins, 226 intracellular signaling cascades cAMP-dependent serine protein kinase A (PKA) activation, 218–219 CCAAT/enhancer binding proteins alpha and beta, 219–221 Gαq/11/PKC pathway, 219 in granulosa cells, 219

intracellular signaling pathways, 218–219, 219f LH surge, 217–218, 218f nitric oxide synthase, 306 proapoptotic and antiapoptotic factors, 306–307 progesterone and prostaglandin (PG) signaling pathways, 306 progesterone receptor, 221 progesterone signaling pathways, 306 proteolytic and angiogenic tissue remodeling angiogenesis, 224 protease actions, 223–224 vasoconstriction and muscle contraction, 224–225 Oxidative stress, 330–331

P

p53, 5–6, 558–559, 577–578, 582 Paclitaxel, 547, 554, 560, 563, 565 Palbociclib, 560 Paracrine, 8 Paramesonephric ducts. See M€ ullerian ducts PARP inhibitors. See Poly-ADP-ribose polymerase (PARP) inhibitors PAX2, 512, 516 Pax2, 532–534, 536 PAX8, 512, 513f, 516 Pazopanib, 563 p120-catenin, 530 PCOS. See Polycystic ovary syndrome (PCOS) PDGF receptor family (PDGFR), 560–561 Pegylated liposomal doxorubicin, 554, 558–559, 563–565 Pelvic radiotherapy, 493 Pembrolizumab, 561, 586 Pentraxin 3 (PTX3), 117 Peptides, 5 Perfluoroalkyl acids (PFAAs), 446–447, 486 Perfluorooctanesulfonate, 486 Perfluorooctane sulfonic acid (PFOS), 446–447 Perfluorooctanoic acid (PFOA), 446–447 Perforin, 583–584 Perimenopause, 99 Permethrin, 151 Perrault syndrome, 473 Personalized cancer therapy (PCT), 588 Pertuzumab, 557–558 PGCs. See Primordial germ cells (PGCs) PGF2A. See Prostaglandin F2A (PGF2A) Pgr gene (PRKO), 221 Phosphatase and tension homolog (PTEN), 29, 296, 581 Phosphatidylinositol-4,5-bisphosphate 3kinase, 296 Phosphatidylinositol 3,4,5-triphosphate (PIP3), 29 Phosphoinositide 3-kinase (PI3K) signaling pathway, 6, 6f Phosphomannomutase deficiency, 476 Phthalates, 446, 486–487 Phytoalexin resveratrol (RSV), 587–588 PI3K/AKT signaling pathway, 548, 555f aberrations within, 555

607 description, 581 downstream effector, 555 granulosa cell differentiation and proliferation by, 40 inhibitors, 556 and MEK inhibitor combination therapies, 565 in ovarian cancer chemoresistance, 581–582 primordial follicle activation, 29–32, 30f PIK3CA mutations, 517 PI3K/PTEN signaling, 517 Pilaralisib (GDC-094), 556 Pilosebaceous unit (PSU), 418–419 Pioglitazone, 424 PKD signaling pathways, 108 p27KipI, 27 Placental aromatase activity, 445 Plasma gelsolin (pGSN), 585 Platelet-derived growth factor (PDGF), 560–561 Platinum-based chemotherapy, 540, 551–552, 554, 563–564 Platinum-taxane therapy, 563 Pleiotropic mendelian disorders, 474 Pluripotency transcription factors, 299 PND23, 24 POF, 1B (POF1B), 465 Poly-ADP-ribose polymerase (PARP) inhibitors, 548, 562 BRCA-mutated tumors, 549 categories, 549 clinical trials, 549, 550t and DNA damage repair, 549, 550f niraparib, 552 olaparib, 551 rucaparib, 551–552 side effects, 549 talazoparib, 553 veliparib, 552–553 Polyaromatic hydrocarbons (PAHs), 486 Polycystic ovarian morphology (PCOM), 418 Polycystic ovary syndrome (PCOS), 41–42, 405–406, 485, 536 activin and follistatin, 101 advanced glycation end products, 452–454, 454f AMH, 119, 408 androgen-secreting tumors, 416 antimullerian hormone, 421 characteristics, 311, 437 chronic anovulation, 419 clinical manifestations, 311 clinical sequelae of cardiovascular disease, 422–423 gynecologic cancer, 421–422 infertility and chronic anovulation, 421 treatment of, 423 type 2 diabetes mellitus, 422 Cushing’s syndrome, 416–417 diagnosis and clinical stigmata, 417 dietary-related weight gain, 447–449 differential diagnosis of, 415, 416t environmental factors, 443 environmental toxins

608 Polycystic ovary syndrome (PCOS) (Continued) Bisphenol A, 446 endocrine-disrupting chemicals, 445–446 nicotine, 447 perfluoroalkyl acids, 446–447 phthalates, 446 triclocarban, 446 epigenetic modifications definition, 441 differential methylation, 442 hypermethylation, 441 hypomethylation, 442 micro-RNA, 442–443 X-chromosome inactivation, 442 fetal programming hypothesis animal-based studies, 443–444 human observational studies, 444–445 mechanisms, 443 generalized treatment strategies of lifestyle modification, 423 metformin, 424 ovarian suppressive therapies, 424 thiazolidinediones, 424 genetic and epigenetic features, 443, 444f etiology, 428 genetic factors candidate gene studies, 439–441 and environmental factors, 149–150 genome-wide association studies, 438–439 oligogenic condition, 438 genome-wide association studies, 428–429 across ethnicities, 318 in European populations, 317–318 in Han Chinese population, 311–317 in Korean populations, 317 list, 312–316t gut microbiota, 449–450 hemoglobin A1C, 420–421 hirsutismis, 418–419, 424–425 hyperandrogenemia, 418 hyperandrogenism, 418 hyperprolactinemia, 415–416 hyperthecosis, 417 hypothyroidism, 415 identification of, 417–418 infertility, treatment of clomiphene citrate, 425–426 gonadotropins, 426 in vitro fertilization, 426–427 laparoscopic ovarian drilling, 426 letrozole, 426 lifestyle modification, 425 inherent problems, 437–438 inhibin and, 101 insulin resistance, 419–420 lack of exercise and physical activity cardiovascular health, 450–451 physical fitness, 450 psychological well-being, 451 reproductive function, 450 lifetime disorder, 427–428 metformin administration, 119 next-generation sequencing, 428–429 nonclassical congenital adrenal hyperplasia, 416

INDEX

oral glucose tolerance testing, 420–421 phenotypic variation, 428 prevalence, 149–150, 311 socioeconomic status, 451–452 symptoms, 149–150 TGF-β superfamily, 119 Positive regulatory domain 1 (Prdm1), 296 Positive regulatory (PR) domain, 296 Post-GnRHa trigger LPS with hCG, 369–372, 370–373f intense, 369, 370f with LH, 373, 374f mid-luteal phase, 369 nasal buserelin three times a day, 373–374, 374f to ovarian responses, 374 luteolysis, 367–368 Preadipocytes, 521 PRECEDENT trial, 558 Prednisolone, 503 Pregnancy activin A, 100 clinical, 388, 389f corpus luteum (see Corpus luteum (CL)) follistatin, 100 inhibin A, 100 luteolysis (see Luteolysis) rates, 224, 351–352, 392 Pregnancyassociated ovarian hyperstimulation syndrome (pOHSS), 140–141, 144 Preimplantation genetic diagnosis (PGD), 327 Preimplantation genetic screening (PGS) hypothesis, 390–391 Preimplantation genetic testing for aneuploidy (PGT-A), 391 Premature ovarian failure (POF), 461, 478 autosomal-dominant, 479 polygenic factors in, 480 sporadic and familial patients, 466–471t Premature ovarian failure/insufficiency (POF/I) characteristics, 318–319 complication, 318–319 cytogenomic studies of, 321–322 female infertility, 318–319 genetic risk factors, 319 GWAS across ethnicity, 319–321 in Dutch population, 319 Han Chinese population, 319 list, 320t prevalence, 318–319 symptoms, 318–319 Premature pubarche, 427 PRIMA, 552 Primary amenorrhea, 502 Primary ovarian failure (POF), 381, 464, 474 Primary ovarian insufficiency (POI), 381. See also Premature ovarian failure (POF) AMH level, 29 associated BMP15 mutations, 36, 118–119 characteristics, 150 GDF9 gene mutation, 118–119 inhibin B and, 99–100

mechanism, 150 NOBOX mutation, 118–119 Primary tumors, 519 Primordial follicle activation intracellular signaling pathways granulosa cell differentiation, 33–34 mTOR signaling, contribution of, 32–33 oocyte activation, regulation of, 29–32, 31f oocyte-specific transcriptional networks, 33 regulation of, 29–34 Primordial follicle formation Figla regulation, 303 germ cell nest breakdown and, 24–29 growth factors and somatic cell cycle progression, 27–28 neonatal oocyte survival, molecular control of, 25–26 notch signaling, regulation, 28–29 steroid hormones, 26–27, 26f Notch signaling pathway, 302 occurrence, 206 region of, 73–74t Primordial follicles, 159–160, 487, 504 activation of, 6–7 death, selection for, 5–6 density, 495 dynamic changes in, 7–8 ECM in, 498 ovarian reserve of, 5 ovarian tissue cryopreservation, 493 puberty, 485, 493 Primordial germ cells (PGCs), 23–25, 112, 157, 161, 165, 461 differentiation and development, 205 gonadal ridge formation, 75 proliferation and migration, 205–206 survival and migration, 299 Proangiogenic factors, 522 Procarbazine, 503 Progesterone, 29 Progesterone production AMP-activated protein kinase activation, by PGF2α, 243–244 activation loop, 242–243 AMP:ATP/ADP:ATP ratios, 242–243 inactivation, 243 inhibition, by LH, 243 pharmacological activators, 242–243 phosphorylation, 243 to restore energy homeostasis, 242–243 cholesterol storage, in lipid droplets, 239–241, 240f corpus luteum extensive vascular network, 238 functional lifespan, 238 immune cell infiltration, 238 LH, 239 luteal regression, 238 menstrual or estrous cycle, 238 gene regulation cytochrome P450 side-chain cleavage (P450scc) complex, 245 HMGCR, 247–248 HSD3B1/2, 246 LDLR, 247 LIPE, 248

INDEX

NCEH1, 248 SCARB1, 247 STAR, 244–245 hormone-sensitive lipase, 241–242 Progesterone receptor, 221 Progesterone receptor membrane component 1 (PGRMC1), 465 Progesterone signaling pathways, 306 Progestin-only pills, 506 Programmed cell death 1 (PD-1), 561, 586 Programmed cell death ligand 1 (PD-L1), 561–562, 584, 586 Programmed cell death ligand 2 (PD-L2), 586 Progression-free survival (PFS), 551–556, 563–564, 581 Prolactin, 415–416 Proliferation of germ cells (POG), 474 Prophase arrest, 165–166 cAMP, oocyte promotion, 168–169 cGMP, 169 Prostaglandin (PG), 226 Prostaglandin endoperoxide 2 (PGE2) luteal angiogenesis, 261 matrix remodeling, 261–262 Prostaglandin-endoperoxide synthase 2 (PTGS2), 114–115 Prostaglandin F2A (PGF2A) actions on endothelial cells, 274–275 actions on steroidogenic cells ATF3, 273 downstream effects, 273, 274f induced progesterone production, 272–274 in vitro studies, 272 mechanism of action, 273 luteolytic capacity, 270–271 Prostaglandin F receptor (PTGFR) expression, 270 Prostaglandin (PG) signaling pathways, 306 Prostaglandin synthase 2 (PGS-2), 306 Protein kinase B (PKB), 581 Protein phosphatase magnesium-dependent 1 D (PPMID) expression, 578 Protein phosphatases (PPs), 184–185, 185f Prsistent M€ ullerian duct syndrome (PMDS), 403 PTEN. See Phosphatase and tension homolog (PTEN) PTEN/PI3K signaling, 206–207 Puberty, 100

Q

QUADRA, 552 Quantitative polymerase chain reaction (qPCR), 329 Quorum-sensing model, 27

R

R04929097, 559 Radiotherapy, 493 Ramucirumab (IMC-1121B), 564 Ras homolog enriched in brain-GTP (RhebGTP), 555 RAS/RAF pathway, 548, 556, 556f Reactive oxygen species (ROS), 225–226 increased production, 331–332 induce apoptosis, 331–332 mitochondria respiratory chain, 331

Receptor for advanced glycation end products (RAGE, 453 Receptor tyrosine kinase (RTK), 27 Reduced representation bisulfite sequencing (RRBS), 211–212 Regulatory-associated protein of mTORC1 (RPTOR), 32 Reproductive toxicants, 485 Response evaluation criteria in solid tumors (RECIST), 551–552 Retinoic acid (RA), 23 Retinoic acid gene 8 (Stra8), 23 Revascularization process, 497–498 Ribonucleoprotein particles (RNPs), 209 Ribosomal protein S6 (rpS6), 32 Ribosomal s6 inase 1 (S6K1), 32 RNA binding, 205–206 RNA-Seq, 530 ROS. See Reactive oxygen species (ROS) Rosiglitazone, 424 Rotterdam criteria, 415, 417–418 R-spondin1 (Rspo1), 298 Rucaparib (CO-338), 551–552

S

Saikosaponin-d (Ssd), 587–588 Saracatinib (AZD0530), 560 SART. See Society for Assisted Reproductive Technology (SART) Scavenger receptor-type B1 (SCARB1), 238f, 239, 247 Secondary amenorrhea, 502 Second embryonic kidney, 512 Secreted frizzled-related protein 2 (SFRP2), 522, 523f Secretory gelsolin (sGSN), 576 Selectable follicles, 51 Selection, unified theory of, 55–56 Selumetinib, 556 Seribantumab, 558 Serous tubal intraepithelial carcinomas (STICs), 516 Sertoli-cell-only syndrome, 474 Sertoli cells, 403 Sertoli-leydig cell tumor (SLCT), 416, 518 Sex cord-stromal ovarian tumors, 514 adult granulosa cell tumors, 518 sertoli-leydig cell tumors, 518 Sex hormone-bindings globulin (SHBG), 418–420, 444–445 Sex steroid hormones, 5 Sf1-cre mouse model, 299 SHBG. See Sex hormone-bindings globulin (SHBG) SH2 domain-containing inositol phosphatase (SHIP), 581 Signal transducer and activator of transcription 1 (STAT1), 520 Simvastatin, 498 Single-cell RNA-seq techniques, 208–209 Single-nucleotide polymorphisms (SNPs), 311, 439 Slit/Robo pathway functions, 303–304 Slow freezing, ovarian tissue cryopreservation, 493–494 SMAD-dependent signaling pathway, 108 SMAD4 mutations, 524

609 Small cell carcinoma of the ovary, hypercalcemic type (SCCOHT), 559–560 Small-for-gestational age newborns, 445 SMARCA2 protein, 548 SMARCA4 protein, 548 Smith-Lemli-Opitz syndrome, 86 Smooth muscle activating endothelins, 224–225 SNAI1, 537–538 Snail transcription factors, 539 Society for Assisted Reproductive Technology (SART), 384, 408 Socioeconomic status (SES), 451–452 SOLO-2 studies, 551–552 Soluble receptor for advanced glycation end products (sRAGE), 453 Somatic cells, 52–53 Sorafenib, 564 Soy phytoestrogen genistein, 26–27 Spermatogenesis and oogenesis specific basic helix-loop-helix 1 (Sohlh1), 33, 199, 303, 303t, 465–466 Spermatogenesis and oogenesis specific basic helix-loop-helix 2 (Sohlh2), 33, 199, 303, 303t Sphingosine-1-phosphate (S1P), 498, 502 Spironolactone, 425 Spontaneous OHSS, 348 Sporadic and familial patients with POF, 466–471t SRC-family kinases, 560 Stable disease, 562–563 STAG3 (POF8), 472 STAT3 signaling pathway, 521–522 Stem-A, 536–537 Stem-B, 536–537 Stem Cell Antigen 1 (Sca1/Ly6a), 532 Stem cell-based techniques, 504 Stemness epithelial-to-mesenchymal transition, 531–532 in fallopian tube epithelium, 532–534, 533t, 534f in ovarian surface epithelium identification and characterization, 532 regulation of, 532, 533t Stemness and epithelial-to-mesenchymal transition, 531–532 Steroid hormones, 115 biosynthesis intracellular concentrations of free radicals, 278 lipoproteins for, 273 and metabolism, 88–91 granulosa cell proliferation, 40–42 role in nest breakdown, 26–27, 26f Steroid metabolism, 448 Steroidogenesis, 487–488 3β-HSD/Δ5-4, 91 cytochrome P450 cholesterol side chain cleavage enzyme and, 90–91 defects in cholesterol, 87 de novo, 85–86 descriptions, 88 expression of genes, regulation of GATA4 and GATA6, 92

610 Steroidogenesis (Continued) genes encoding proteins, 92 lipases, 86–87 lipoproteins, 85–86 steroidogenic acute regulatory protein, 88–90 TSPO stimulate, 89–90 Steroidogenic acute regulatory protein (STAR), 237–238, 244–245 Steroidogenic cells ATF3, 273 downstream effects, 273, 274f induced progesterone production, 272–274 in vitro studies, 272 mechanism of action, 273 Steroidogenic factor 1 (SF1), 472 Steroidogenic luteal cell, 238f Steroid production and metabolism, 440 Stimulated by retinoic acid gene 8 (STRA8), 23, 206, 296 STRAW (Stages of Reproductive Aging Workshop), 99 Structural molecule activity, 205–206 Subcortical Maternal Complex (SCMC), 209 Sunitinib, 564–565 Surrounded nucleolus (SN), 173–174 Survivin, 578–579 SWITCH/sucrose nonfermentable (SWI/SNF) chromatin-remodeling complex, 548, 559–560 SYCE1 (POF12), 472–473 Synaptonemal complex protein 3 (SYCP3), 298–299

T

TAK1, 581 Talazoparib (BMN673), 549, 553 Tamoxifen, 502, 557 Tamoxifen citrate, 565 TAp63, 5–6 Targeting epigenetic changes, 587 TATA-binding protein 2 (TBP2), 207 TATA-binding protein-associated factor 4B (TAF4B), 207, 300–301, 320–321 Tellurium-based immunomodulatory agent, 502 Temsirolimus, 555–556, 565 Testosterone, 10, 41, 418, 444, 449 Tex14, 24–25 TGFβ. See Transforming growth factor β (TGFβ Theca cells, 38 The Cancer Genome Atlas (TCGA), 547–548 Thiazolidinediones, 424 Three-parent embryos, 338 Thrombocytopenia, 552–553 Thyroid-stimulating hormone (TSH), 415 Thyrotropinreleasing hormone (TRH), 415 TILs. See Tumor-infiltrating lymphocytes (TILs) Time-to-pregnancy (TTP), 486 Tissue inhibitor of matrix metalloproteinases (TIMPs), 262 Tissue-specific mitochondria, 328 Tissue-type plasminogen activator (tPA), 262 TME. See Tumor microenvironment (TME) Topotecan, topotecan, 554, 557–558, 563

INDEX

TP53 mutations, 514–516, 547–548 Trametinib, 565 Transcelomic dissemination, 520 Transcription factors, epithelial-tomesenchymal transition, 530–531 Transforming growth factor β (TGFβ), 107, 296, 487, 518, 539–540 receptors in human ovary, 111–112, 111t type I and type II, 108 TGFB1, 532, 535 TGFβP3, 112 Transforming growth factor β superfamily members, 95–96 activins/inhibins, 107 anti-M€ ullerian hormone, 29, 107 bone morphogenetic proteins, 107 cumulus-oophorus complex, 116–117 ECM modulation, 116 in female reproductive pathology Bmp4/Bmp7 gene mutation, 118 endometriosis, 119 GDF9 and BMP15, 118 knockout mice model, 118 mouse models, 118 PCOS, 119 primary ovarian insufficiency, 118–119 GC proliferation and terminal differentiation, 116 GDFs, 107 in human ovary, 114f intraovarian cell-cell communication, 113 ligands BMP15, 35–36 homodimers/heterodimers, 107–108 in human ovary, 109–111, 109–110t subfamilies, 107–108 luteal function, 117–118 oocyte-somatic cell interactions, 113–115 ovarian preantral follicle development, 112–113 ovarian steroidogenesis, 115–116 ovulation, 117 pathophysiological roles, 119–120 PGCs development, 112 signal transduction in ovary, 35–37 pathway, 108 therapeutic development, 119–120 Transgenerational epigenetic inheritance, 152t BPA, DBP and DEHP, 152 DDT, 152 definition, 149 dioxin, 152–153 epigenetic information requirement, 149 exposure to toxicant, 149 factors, 150–151 JP-8 (hydrocarbon), 153 methoxychlor, 152 mice model, 149, 150f permethrin and DEET, 151 vinclozolin, 151 Transgenic mouse models antral and preovulatory follicle development, 303–305 folliculogenesis

Fanconi anemia family, 303 Gas2, 302 GCC breakdown, 300–301 germ cell deletion, 302 Hes1, 302 Hippo signaling pathway, 302–303 meiotic arrest, 305–306, 305f mTORC, 301–302 Notch signaling pathway, 302 oocytes, loss of, 300–301 oocyte-specific transcription factors, 303, 303t PDK1-Kit-PI3K/PTEN/AKT-FOXO3 signaling network, 301 PI3K/AKT/mTORC1 signaling, 301, 301f primordial follicles formation, 300–301 secondary follicle stage, 303 TAF4B, 300–301, 301f TSC1 and TSC2, 301–302 germ cell development, 296–300 histology, 296, 297f meiosis resumption, 305–306 ovarian differentiation, 299 ovulation, 306–307 Translation initiation, 205–206 Translocator protein (TSPO), 89–90 Transzonal projections (TZPs), 28 Trastuzumab, 557–558 Trebananib (AMG 386), 563 Tremelimumab, 561, 586 3,4,40-Trichlorocarbanilide, 446 Triclocarban (TCC), 446 Triplet nucleotide repeats (CGG), 478, 478f Triplet repeats (CTG), 479 Troglitazone, 424 Tuberous clerosus complex 1 (Tsc1), 207 Tuberous sclerosis complex 1 (TSC1), 301–302 Tuberous sclerosis complex 2 (TSC2), 301–302, 555 Tumor-associated macrophages (TAMs), 583 Tumor-derived extracellular vesicles, 584–585 Tumor-Infiltrating APCs, 583 Tumor-infiltrating lymphocytes (TILs), 562, 583–584 Tumor microenvironment (TME) chemoresistance, 575–576, 577f, 582–584 cancer-derived extracellular vesicles, 585 cellular mechanisms, 576 cytokines/chemokines, 586 gelsolin, 576–577, 578–580f, 582 hexokinase II, 581–582 inhibitor of apoptosis proteins, 577–578 P53, 577–578, 582 PI3K/Akt pathway, 581–582 plasma gelsolin, 585 tumor-derived extracellular vesicles, 584–585 ovarian cancers, 519 angiogenesis and neovasculature, 522–524 cancer-associated fibroblasts, 520 in disease progression, 520–525, 525f mesenchymal stem cells, 520–521 ovary-associated adipocytes, 521–522

611

INDEX

Tumor necrosis factor (TNF), 535 Tumor necrosis factor superfamily-15 (TNFSF15), 522 Turner syndrome, 462, 493 Twist transcription factors, 539 Two-cell two gonadotropin theory, 9 Type 2 diabetes mellitus, 422, 427 Type I error, risk of, 486–487

U

Ultrasonography, 51, 58–61 Undifferentiated carcinomas, 514 Unopposed estrogen, 421–422 Urokinase-plasminogen activator (uPA), 262

V

Vandetanib, 564 Variable number of tandem repeats (VNTR) region, 440 Vascular endothelial growth factor (VEGF), 487 and angiogenesis, 553–555 bevacizumab, 553 biological actions, 346 in COS patients, 346 gene, 346 graft revascularization, 498 growth factors, family of, 553, 554f hCG mediator, 346 in OHSS, 346 proangiogenic factors, 583 Vascular endothelial growth factor A (VEGFA), 257, 260–261 Vascular endothelial growth factor receptor 3 (VEGFR3) signaling, 79 Vascular epithelial growth factor (VEGF), 522 Vascular permeability factor (VPF), 346 Vasculature system, 583 VDAC. See Voltagedependent anion channel (VDAC) Veliparib (ABT-888), 549, 552 Verapamil, 532 Vinclozolin, 151 Vincristine, 503 Vintafolide, 558 Vitamin D deficiency, 405–406

Vitamin E, 498 Voltagedependent anion channel (VDAC), 577–578, 581 VO2 max, 450 Vorinostat, 587 V-RAF 1 murine leukemia viral oncogene homolog 1 pathway, 556

W

WC1+ gamma delta T cells, 276 Weight loss, 423, 448–449 Whole genome bisulfite sequencing (WGBS), 211–212 Wild-type and mutant mouse models, histology of, 297f Wilms’ tumor-1 (WT1) gene, 299, 474–476 Wingless-type MMTV integration site (WNT), 296 Wingless-type MMTV integration site family, member 4 (WNT4), 298 WNT16B, 522, 523f Wnt/β-catenin/Tcf (Wnt) signaling, 517 Wolffian ducts, 512, 513f Wolman’s disease, 84–85 Wound repair, phenotypic plasticity in, 534–536 cell plasticity, potential consequences of, 536 inflammation, 535 reepithelialization, 535 WT1 (Wilms’ tumor-1) gene, 299, 474–476 WT1 vaccination, 561

X

X chromosome, haploinsufficiency for, 462–465 candidate genes for Xp gonadal determinants, 464 Xq gonadal determinants, 464–465 deletions X long-arm, 464 X short-arm, 462–463, 463f monosomy X, 462 nomenclature, 462, 462t X-chromosome inactivation (XCI), 442 Xenograft models, 494, 498

XIAP. See X-linked inhibitor of apoptosis protein (XIAP) X-inactivation center (XIC), 464 X-linked inhibitor of apoptosis protein (XIAP), 578–581 X-linked zinc finger protein (ZFX), 299 X long-arm deletions (Xq), 464 Xp gonadal determinants, candidate genes for, 464 XPNPEP2, 465 Xq gonadal determinants, candidate genes for, 464–465 X short-arm deletions, 462–463, 463f XX gonadal dysgenesis cerebellar ataxia, 473 clinical spectrum of, 465t FSHR gene mutation (C566T), 480–481 in malformation syndromes, 474, 474t perrault syndrome, 473 without somatic anomalies, 465–473, 466–471t CSB-PGBD3 (POF11), 472 FIGLA (POF6), 472 HFM1 (POF9), 472 MCM8 (POF10), 472 MSH5 (POF13), 473 NOBOX (POF5), 466–471 NR5A1 (POF7), 472 sporadic and familial patients with POF, 466–471t STAG3 (POF8), 472 SYCE1 (POF12), 472–473 46,XY Agonadia, 477–478 46,XY Disorders of sex development (DSD), 472, 475–476 Xyotax, 588

Y

Y chromosome, 461 Yolk-sac, 157

Z

Zona pellucida (ZP), 4, 23–24 Zuclomiphene, 425–426 Zygotic arrest-1 (Zar1), 158

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