Chromosomal Complements in Human Immature and Mature Oocytes in Relation to Maternal Age

Wejdan M. Alenezi

Department of Human Genetics McGill University Montreal, Quebec, Canada

Submitted March 2015

A thesis submitted to McGill University Faculty of Graduate Studies and Research in partial fulfillment of the requirements for the degree of masters in science

©Wejdan M. Alenezi

i ACKNOWLEDGMENT

Foremost, I dedicate my M.Sc. thesis to my beloved mother and siblings whose overseas, unlimited love and support pushed me to go further on the way of knowledge. Their encouragements and motivations allowed me to undertake this academic study and made my

M.Sc. thesis to be possible achieved.

I would like to express my sincere gratitude to my supervisor Prof. Asangla Ao for introducing me to the field of preimplantation genetic diagnosis (PGD) and embryology. She took the initiative to teach me the basics of human embryo manipulation and the fundamentals of

PGD practice. She generously offered me her valuable time to guide me throughout writing my reports and M.Sc. thesis. She encouraged me to participate in several PGD-related conferences.

On the top, she emotionally supported me through my health issues I have had during my studies. Without her unwavering guidance, supervision and patience, this academic accomplishment would not have been possible.

Beside my supervisor, I would like to extend my acknowledgments to Dr. Zhange Li and

Xiao Yun Zhang for teaching me most important PGD related techniques. Their helpful guidance, endless patience, and encouraging words made this project achieved.

I am also indebted to my supervisory committee members Prof. Anna Naumova and Prof.

Teruko Taketo for their insightful comments and suggestions, and valuable assistance and guidance (especially in statistics) that made this project to be successfully accomplished.

I extend my acknowledgments to all embryologists, nurses, and physicians at McGill

Reproductive Center for their assistance and cooperation during these years. I am deeply grateful to all patients who donated the research samples. I would like also to acknowledge the

ii scholarship granted from the Department of Applied Medical Sciences at Taibah University, my hometown university.

Finally, I would like to express my heartfelt thanks to my long-suffering friends: Maha,

Raha, Basma, Ghadeer, Luci and Amal whose endless emotional support allowed me to overcome all obstacles I faced throughout my academic studies. Also, I am deeply thankful to

Prof. Idrees Al-Turk who encouraged and motivated me to pursue my graduate studies. His continues support and advice, as well as absolute faith in me allowed me to undertake all difficulties I faced throughout my studies.

iii PRESENTATION OF THE CURRENT THESIS

This M.Sc. thesis was prepared in accordance to the guidelines for the traditional, monographic thesis style. The table of content, and an abstract in English and in French are included. Chapter one contains a general introduction, which provides an overview of the research scope of the present project including the significance, the rationale, and the objectives of the M.Sc. project. This is followed by a literature review, which provides a comprehensive review of the relevant literatures. Chapter two includes the material and methods were applied in the present research. Chapter three demonstrates the results, followed by chapter four that discuss the results obtained in this project. Last, a summery of the present work and final conclusions, including the future direction are presented in chapter five. This thesis is completed by a list of the references cited within chapter one to five, followed by an appendix includes a list of ethics approval of using all human materials were used in this project.

iv ABBREVIATIONS

0PN Zero Pronuclear Zygote 1PN One Pronuclear Zygote 2PN Two Pronuclear Zygote 3PN Three Pronuclear Zygote AI Anaphase I stage aCGH Array Comparative Genomic Hybridization AL Anaphase Lag AFC Antral Follicular Count CGH Comparative Genomic Hybridization Chr Chromosome Cht Chromatid COH Controlled Ovarian Hyperstimulation FISH Fluorescence In Situ Hybridization GV Germinal Vesicle GVBD Germinal Vesicle Break Down ICSI Intracytoplasmic Sperm Injection IVF In Vitro Fertilization IVM In Vitro Maturation MC Miscarriage MF Male Factor MI Metaphase I Stage MII Metaphase II Stage ND Non-Disjunction PCOS Polycystic Ovary Syndrome PD Premature Separation PGD Preimplantation Genetic Diagnosis PRL Repeated Pregnancy Loss PSSC Premature Separation of Sister Chromatids SGD Single Gene Defects SNP Single Nucleotide Polymorphism Array

v ABSTRACT

Numerical chromosome abnormalities significantly contribute to the high incidence of spontaneous abortions, stillbirths, and live births with congenital defects. The vast majority of these abnormalities are maternally related and attributed to meiotic errors occurring during oogenesis. The clinical relevance of these errors, the contribution to the overall rate of chromosome abnormalities, and the association with advanced maternal age are well documented. Maternally related chromosome abnormalities can also be attributed to errors occurring prior to meiosis. Select studies have estimated the incidence of pre-meiotic mitotic errors, though the prevalence, clinical relevance, and association with maternal age remain unknown. Therefore, we aimed to investigate the incidence of pre-meiotic mitotic errors in human immature and mature oocytes derived from women who underwent controlled ovarian hyperstimulation (COH) treatment cycles in relation to maternal age. FISH-based analysis for chromosomes most frequently involved in pregnancy loss (13, 15, 16, 18, 21, 22, and X) was applied to analyze the chromosomal complements in human oocytes. We pioneered a method in developing and validating the FISH signals scoring criteria of chromosomally normal immature oocytes at different maturation stages (in relation to maternal age) by plotting histograms of

FISH signals per tested chromosome. These distributions allowed us to determine the possibility of sample variation, as well as confirmed our FISH signals scoring criteria for chromosomally normal oocytes. The overall rates of chromosomally normal immature oocytes at all different maturation stages were comparable between both maternal age groups, suggesting that pre- meiotic mitotic errors are not maternal-age dependent. We re-analyzed the data to estimate the incidence of chromosomally normal immature oocytes at the different maturation stages (in relation to maternal age) when each patient donated “sibling oocytes” for eliminating patient

vi bias. These results confirmed that pre-meiotic mitotic errors are not associated with maternal age. Our findings indicate that the incidence of pre-meiotic mitotic errors in human mature oocytes with the corresponding first polar bodies in relation to maternal age is considerably low compared to first meiotic errors. As previously reported, the overall rate of first meiotic errors is directly related to maternal age.

Our data suggest that although the contribution of pre-meiotic mitotic errors to the overall rate of maternally related chromosome abnormalities is relatively low, consequences of these errors have clinical relevance to human fertility, at least in our study population.

vii SOMMAIRE

Plusieurs anomalies chromosomiques contribuent significativement à l’incidence élevée d’avortements spontanés, de morts à la naissance et de naissances avec défauts congénitaux. La vaste majorité de ces anomalies sont attribuées à des erreurs méiotiques qui prennent place durant l’oogenèse. La pertinence de ces erreurs à la pratique clinique, leur contribution aux taux d’anomalies chromosomiques et leur association avec l’âge maternel avancé sont tous bien documentés. Les anomalies chromosomiques d’origine maternelle peuvent aussi être attribuées aux erreurs qui précèdent la méiose. Quelques études ont tenté d’estimer l’incidence d’erreurs miotiques précédant la méiose, malgré que leur prévalence, leur importance clinique et leur association avec l’âge maternel demeurent inconnus. Par conséquent, nous avons investigué l’incidence d’erreurs mitotiques précédant la méiose en relation à l’âge maternel dans les oocytes humains immatures et matures dérivés de femmes ayant subi des cycles de traitement de stimulation ovarienne. La méthode d’analyse FISH pour les chromosomes les plus souvent impliqués dans la perte de grossesse (13, 15, 16, 18, 21, 22, and X) a été appliqué à l’analyse des compléments chromosomiques des oocytes humains. Nous avons créé une nouvelle méthode pour le développement et la validation des critères de notation pour les signaux FISH provenant des oocytes immatures avec chromosomes normaux à différentes étapes de leur maturation en relation avec l’âge maternel. Pour ce faire, nous produisons des histogrammes des signaux FISH pour chaque chromosome testé. Ces distributions nous ont permis de déterminer la variation de l’échantillon ainsi que de confirmer nos critères de notation des signaux FISH.. Le nombre total d’oocytes immatures avec des chromosomes normaux à toutes les étapes de maturation était comparable entre les deux groupes d’âge maternel, suggérant que les erreurs mitotiques qui précèdent la méiose ne sont pas dépendantes sur l’âge maternel. Pour éliminer le biais de

viii sélection, nous avons ré-analysé nos données pour estimer l’incidence de chromosomes normaux dans les oocytes immatures en relation à l’âge maternel lorsque chaque patient a donné des

« oocytes sœurs ». Ces résultats ont confirmé que les erreurs mitotiques précédant la méiose ne sont pas associées avec l’âge maternel. Nos trouvailles indiquent qu’en relation avec l’âge maternel, l’incidence des erreurs mitotiques pré-méiotique dans les oocytes humains matures avec les premiers globules polaires correspondants est beaucoup plus basse que les premières erreurs méiotiques. Le taux général de premières erreurs méiotiques est directement relié à l’âge maternel, comme cela a été reporté auparavant.

Nos résultats suggèrent que malgré la contribution des erreurs mitotiques précédant la méiose au taux général d’anomalies chromosomiques d’origine maternelle est relativement basse, les conséquences de ces erreurs ont une pertinence pour la pratique clinique de la fertilité humaine, du moins dans notre population d’étude.

ix TABLE OF CONTENTS

ACKNOWLEDGMENT.………………………………………………………………………..ii ABBREVIATIONS……………………………………………………………………………...iii ABSTRACT……………………………………………………………………………………...iv SOMMAIRE……………………………………………………………………………………...v TABLE OF CONTENTS……………………………………………………………………….vi LIST OF FIGURES…………………………………………………………………………….vii LIST OF TABLES……………………………………………………………………………..viii

CHAPTER ONE: INTRODUCTION AND LITERATURE REVIEW……………………...1 1.1. INTRODUCTION…………………………………………………………………………..2 1.1.1. Overview of Chromosome Abnormalities in Human Immature and Mature Oocyte……..2 1.1.2. The Rationale of the Study………………………………………………………………11 1.1.3. The Objectives of the Project…………………………………………………………….12 1.1.4. The Significance of the Study……………………………………………………………13 1.2. LITERATURE REVIEW…………………………………………………………………14 1.2.1. Characteristics Chromosome Abnormality in Human…………………………………...14 1.2.2. Significance and Incidence of Chromosome Abnormalities in Human Reproduction…..16 1.2.3. Origin of Chromosome Abnormalities in Human……………………………………….24 1.2.3.1. Chromosomal Complement During Human Oogenesis………………………………..24 1.2.3.2. Pre-Meiotic Mitotic Errors……………………………………………………………..35 1.2.3.3. Meiotic Errors…………………………………………………………………………..41 1.2.3.4. Fertilization Errors……………………………………………………………………...46 1.2.3.5. Post-Zygotic Errors……………………………………………………………………..48

CHAPTER TWO: MATERIAL AND METHODS…………………………………………..54 2.1. FISH Analysis Standardization and Validation (Pre-examination Stage)………………….55 2.1.1. FISH Probe Mixtures Preparation……………………………………………………55 2.1.2. FISH Probes Mixtures Validation……………………………………………………56 2.1.2.1. Human Lymphocytes Isolation and Fixation………………………………………..56 2.1.2.2. Spare Human Embryos Manipulation and Fixation…………………………………57 2.1.2.3. FISH Analysis and Signals Scoring…………………………………………………58 2.2. Experimental Study population (Examination Stage)……………………………………..61 2.2.1. Patients Characteristics…………………………………………………….………...61 2.2.2. Controlled Ovarian Stimulation (COH) Treatment Cycles………………………….65 2.2.3. Oocytes Collection…………………………………………………………………..65 2.2.4. Oocytes Morphological Assessment For Chromosomal Complement Analysis……66 2.2.5. Oocytes Fixation and FISH Analysis………………………………………………..72 2.2.6. Oocytes FISH Signals Scoring………………………………………………………72 2.3. Statistical Analysis and Data Interpretation (Post-examination Process)………………...73

CHAPTER THREE: RESULTS………………………………………………………………74 3.1. FISH Technique Validation in Human Lymphocytes and Spare Embryos, and FISH Signals Scoring in Human Immature and Mature Oocytes………………………………..75 3.1.1. FISH Analysis Efficiency in Human Lymphocytes and Spare Embryos……………75

x 3.1.2. FISH Analysis Efficiency in Human Immature and Mature Oocytes……………….75 3.1.3. FISH Signals Scoring in Human Immature and Mature Oocytes……………………78 3.2. FISH Signals Validation in Human Immature Oocytes…………………………………...85 3.2.1. Distributions of FISH Signals in Mature Oocytes and Unfertilized Oocytes as Control Groups………………………………………………………………………85 3.2.2. Distributions of FISH Signals in Immature Oocytes at Different Maturation Stages………………………………………………………………………………..86 3.3. The Estimated Rate of Chromosomally Normal Immature and Mature Oocytes of Total Study Group in Association to Maternal Age……………………………………………..96 3.3.1. Overall Estimated Rate of Chromosomally Normal Immature and Mature Oocytes……………………………………………………………………………...96 3.3.2. Overall Estimated Rate of Chromosomally Normal Immature and Mature Oocytes in Relation to Maternal Age……………………………………………………………96 3.4. The Estimated Rate of Chromosomally Normal Immature and Mature Oocytes of Sibling Group in Association to Maternal Age………………………………………………….....99 3.4.1. Cohort of Sibling Immature and Mature Oocytes…………………………………...99 3.4.2. Overall Estimated Rate of Chromosomally Normal Immature and Mature Oocytes……………………………………………………………………………...99 3.4.3. Overall Estimated Rate of Chromosomally Normal Immature and Mature Oocytes in Relation to Maternal Age…………………………………………………………..100 3.5. The Estimated Rate of Pre-meiotic Mitotic Errors and First Meiotic Errors in Mature Oocytes in Association to Maternal Age…………………………………………………104 3.5.1. Overall Estimated Rate of Pre-meiotic Mitotic Errors…………………………….104 3.5.2. Overall Estimated Rate of First Meiotic Errors……………………………………109

CHAPTER FOUR: DISCUSSION…………………………………………………………...116 4.1. Pre-Experimental Stage: FISH Technique Standardization and Validation using Human Lymphocytes and Spare Donated Embryos……………………….……………………..118 4.2. Experimental Stage: Chromosomal Complement Analysis in Human Immature and Mature Oocytes Using FISH Technique……………………………………………………...…..120 4.3. Estimated Rates of Chromosomally Normal Immature and Mature Oocytes …………...137 4.4. Estimated Rates of Pre-Meiotic Mitotic Errors in Mature Oocytes….…………………..142 4.5. Limitations………………………………………………………………………………..148

CHAPTER FIVE: CONCLUSION AND FUTURE DIRECTION………………………...150 5.1. Conclusion…………………………………………………………………………………151 5.2. The Study Future Direction………………………………………………………………..152

REFERENCES…..…………………………………………………….………………………154

xi LIST OF FIGURES

CHAPTER ONE: INTRODUCTION AND LITERATURE REVIEW Figure 1.1: The proposed hypotheses for the origin of pre-meiotic mitotic errors in human oocytes…………………………………………………………………………………………....9 Figure 1.2: Oogenesis and spermatogenesis in humans……………………………………..….28 Figure 1.3: A timeline of early oogenesis in humans………………………………………..….29 Figure 1.4: The chromosomal behaviour during the prophase stag in oogenesis…………..…...32 Figure 1.5: The chromosomal complements during pre-meiotic mitotic, meiotic and post-zygotic mitotic divisions in human……………………………………………………………...... 33 Figure 1.6: Proposed models for the alignment and segregation of a trisomic immature oocyte… ……………………………………………………………………………………………………39 Figure 1.7: Proposed mechanisms leading to first and/or second meiotic errors in human mature oocytes………………………………………………………………………………...…………44

CHAPTER TWO: MATERIAL AND METHODS Figure 2.1: The percentiles of both maternal age groups per COH treatment cycles…………...63 Figure 2.2: The infertility diagnosis among patients who underwent COH treatment cycles...... 64 Figure 2.3: Schematic diagram of the morphological assessment of collected oocytes………...68 Figure 2.4: Microscopic images of successfully fixed immature and mature oocytes...... 69 Figure 2.5: Graphical representation of total successfully fixed oocytes at different maturation stages including unfertilized oocytes per treatment cycles………………………………………71

CHAPTER THREE: RESULTS Figure 3.1: Expected normal FISH signals for chromosomes (13, 15, 16, 18, 21, 22, and X) from interphase nuclei from human lymphocytes and spare embryos………………………………...76 Figure 3.2: FISH signals in oocytes at different maturation stages……………………………..81 Figure 3.3: Histograms of FISH signals in relation to maternal age per screened chromosome (13, 15, 16, 18, 21, 22, and X) in oocytes at all different maturation stages including unfertilized oocytes (the color coded histograms represents the FISH probes)………………………………90 Figure 3.4: Graphical representation of successfully fixed and analyzed sibling oocytes at all different maturation stages (when a minimum of one immature oocyte at any maturation stage and one mature oocyte were collected per treatment cycle)……………………………………102 Figure 3.5: Two patterns of FISH signals for pre-meiotic mitotic errors in mature oocytes with the corresponding first polar bodies…………………………………………………………….107 Figure 3.6: FISH signals of pre-meiotic mitotic error in mature oocyte with their corresponding first polar body………………………………………………………………………………….108 Figure 3.7: The percentage of total chromosomally abnormal mature oocytes in relation to maternal age………………………………………………………………………………….…111 Figure 3.8: First Meiotic errors among all analyzed chromosomes in mature oocytes between both maternal age groups……………………………………………………………………….114

CHAPTER FOUR: DISCUSSION Figure 1.4: The chromosomal configuration and FISH signals pattern of immature oocytes post GVBD-B stage………………………………………………………………………………….136

xii Figure 2.4: Non-disjunction of the extra chromosome due to pre-meiotic mitotic error in either cell of MII-1PB doublet………………………………………………………………………...144

LIST OF TABLES

CHAPTER ONE: INTRODUCTION AND LITERATURE REVIEW Table 1.1: Summary of the overall rates of meiotic errors in association to maternal age have been investigated using different techniques for analysis………………………………………....4 Table 1.2: Summary of the overall rate of pre-meiotic mitotic errors have been investigated in human foetal ovarian cells and adult oocytes using different techniques for analysis………..…10 Table 1.3: The Chromosomal Complements in Germ Cells and Human Oocytes at Different stage. ……………………………………………………………………………………………………34

CHAPTER TWO: MATERIAL AND METHODS Table 2.1: The total number, mean, median and range of successfully fixed oocytes at different maturation stage per treatment cycles including unfertilized oocytes in relation to maternal...…70

CHAPTER THREE: RESULTS Table 3.1: Total number of collected, fixed, and successfully hybridized using FISH analysis for all chromosomes…………………………………………………………………………………77 Table 3.2: The expected FISH signals pattern in chromosomally normal and abnormal immature and mature oocytes………………………………………………………………………………80 Table 3.3: Different Patterns of Chromosomal Abnormality in Mature Oocytes……………….83 Table 3.4: FISH signals of all tested chromosomes in immature oocytes at all maturation stages assigned as non-chromosomally normal………………………………………………………....84 Table 3.5: The statistical difference between distributions of FISH signals in relation to maternal age per screened chromosomes in oocyte at different maturation stages, including unfertilized oocytes…………………………………………………………………………………………...95 Table 3.6: The estimated rate of a normal chromosomal complement at different maturation stages in association to maternal age…………………………………………………………....98 Table 3.7: Table of successfully fixed and analyzed oocytes at different maturation stages amongst sibling oocytes (when a minimum of one immature oocyte at any maturation stages and one mature oocyte were collected per treatment cycle)………………………………………...101 Table 3.8: The estimated rate of chromosomally normal immature oocytes across different maturation stages and mature oocytes in association to maternal age in sibling oocytes……...103 Table 3.9: Pre-meiotic errors in mature oocytes and their corresponding first polar bodies..…106 Table 3.10: The frequency of mechanisms leading to errors in mature oocytes in relation to maternal age…………………………………………………………………………………….112 Table 3.11: The frequency of chromosomal and chromatid errors in association to maternal age………………………………………………………………………………………………113

xiii

CHAPTER ONE:

INTRODUCTION AND LITERATURE REVIEW

1 1.1. INTRODUCTION

The research scope of the present project is going to be introduced, including the significance, the rationale, and the objectives of the project prior to the literature review.

1.1.1. Overview of Chromosome Abnormalities in Human Immature and Mature Oocytes

Chromosome abnormality is one of the leading factors that negatively affect human reproduction. Numerical chromosome abnormalities including aneuploidy are known to significantly contribute to the high incidence of spontaneous abortions, stillbirths, and live births with congenital defects (Hamerton, Canning, Ray, & Smith, 1975). The prevalence of these abnormalities is approximated to be 0.3% of live birth defects, up to about 4% of stillbirths, and more than 35% of spontaneous abortions and early embryo losses (J. H. Ford, Wilkin, Thomas,

& McCarthy, 1996; T. J. Hassold, 1986; T. J. Hassold & Jacobs, 1984; Plachot et al., 1987;

Spandorfer, Davis, Barmat, Chung, & Rosenwaks, 2004). The overall rate of numerical chromosome abnormalities is estimated to be about 10% of all clinically recognized pregnancies; this rate is well documented to be directly associated with maternal age. Reported data from in utero conceived pregnancies, as well as assisted conceived pregnancies reveals an overall rate of numerical chromosomal abnormalities to be at least 5%, and as high as 25% of all clinically recognized pregnancies (Griffin, Handyside, Penketh, Winston, & Delhanty, 1991; T. J. Hassold

& Jacobs, 1984) as numerous related studies have been reviewed (T. Hassold & Hunt, 2001;

Jones & Lane, 2013; Nagaoka, Hassold, & Hunt, 2012).

Numerical chromosome abnormalities may arise during gametogenesis (oogenesis and spermatogenesis), around the time of fertilization, and during embryogenesis (C. E. Ford &

Hamerton, 1956). The vast majority of these anomalies are of maternal origin (near to 95%), which arise de novo during oogenesis, whereas about only 5% are paternally contributed

2 (Antonarakis, 1991; T. Hassold, Chiu, & Yamane, 1984; P. A. Jacobs, 1992) as related studies have recently reviewed (Chatziparasidou, Christoforidis, Samolada, & Nijs, 2014; Delhanty &

Handyside, 1995; T. Hassold et al., 1996; T. Hassold, Hall, & Hunt, 2007; T. Hassold & Hunt,

2001; Templado, Uroz, & Estop, 2013). Most of maternally related abnormalities occur during the first and/or second meiotic errors, and recent studies have investigated the prevalence and cytogenesis of meiotic errors in human oocytes derived from women who underwent assisted reproductive technology (ART) treatment cycles (Delhanty, 2005; Fragouli, Wells, & Delhanty,

2011; Kuliev & Verlinsky, 2004; Pellestor, Anahory, & Hamamah, 2005). On average, half of normally fertilized oocytes are chromosomally abnormal due to meiotic errors (Kuliev,

Zlatopolsky, Kirillova, Spivakova, & Cieslak Janzen, 2011). The contribution of these errors to the overall rate of chromosome abnormalities is estimated to be about 70-80% (Rabinowitz et al.,

2012; Sills, Li, Frederick, Khoury, & Potter, 2014). The clinical relevance of meiotic errors to human reproduction and fertility, and the association with advanced maternal age has been well documented (T. J. Hassold & Jacobs, 1984; Kuliev et al., 2011). Women in their 20s are at a lesser risk for meiotic errors (only 2 to 3%), as compared to women in their late 30s who have a higher risk (exceeding 30%). The underlying molecular mechanisms of the adverse effect of advanced maternal age upon the increased rate of chromosome abnormalities remains unclear though many related studies have been recently reviewed (Jones & Lane, 2013; Nagaoka et al.,

2012). Many hypotheses have been proposed to elucidate the effect of advanced maternal age upon increased rates of chromosome abnormalities, as reviewed by (Rowsey et al., 2013). The meiotic arrest model is most favourable when considering the association between the depleted cohesins in mature oocytes derived from older women compared to those derived from younger women (Duncan et al., 2012; Garcia-Cruz, Brieno, et al., 2010; Tsutsumi et al., 2014).

3 Table 1.1: Summary of the overall rates of meiotic errors in association to maternal age has been investigated using different techniques for analysis. Table 1.1-A: Karyotype Analysis Studies Average Maternal Age No. of Female No. of Successfully Type of Oocytes Overall Aneuploidy Rate of oocytes in % (No.) 1 Years (Range) (Cycles) Analyzed oocytes Analyzed (Angell, Aitken, van Look, Lumsden, & Templeton, 1983) 35.1 (31-39) 7 3 Unfertilized 66.6 (2/3) (Martin et al., 1986) 29.4 (24-35) 33 50 Unfertilized 32 (16/50) (Bongso, Chye, Ratnam, Sathananthan, & Wong, 1988) (27-42) 143 251 Unfertilized 21.1 (53/251) (Djalali, Rosenbusch, Wolf, & Sterzik, 1988) 31 (24-39) 62 96 Unfertilized 30.2 (29/96) (Pellestor & Sele, 1988) 30.6 (22-40) 87 201 Unfertilized 18.6 (35/188) (Plachot et al., 1988) DNA2 117 296 Unfertilized 24.1 (55/228) and 38.3 (26/68)3 (Delhanty & Penketh, 1990) DNA2 DNA2 75 Unfertilized 3.8 (2/53) and 9.1 (2/22) (Gras, McBain, Trounson, & Kola, 1992) DNA2 56 88 Unfertilized 34.5 (20/58) and 23.3 (7/30) (Benkhalifa, Menezo, Janny, Pouly, & Qumsiyeh, 1996) (21-42) 208 135 Unfertilized 19 (45/135) (Roberts & O'Neill, 1995) 33.5 (23-42) DNA 233 Unfertilized 22.3 (52/233)3 (Macas, Imthurn, Roselli, & Keller, 1996) 33.9 (20-38) 48 (48) 28 Unfertilized 32.1 (9/28) (Pellestor, Andreo, Arnal, Humeau, & Demaille, 2003) 33.7 (19-46) 792 1397 Unfertilized 20.1 (280/1397)3 1 The chromosomal complements were investigated in only unfertilized oocytes, but not polar bodies. 2 DNA refers to “data not available” which were not mentioned by the author in the study. 3 The overall rates of aneuploidy showed a statistical difference in relation to maternal age (two groups of women were 35 years old or younger, and women were older than 35 years)

4 Table 1.1-B: FISH Analysis Studies Average No. of FISH Probes No. of Analyzed Oocytes Type of Oocytes Overall Rate of Aneuploid Cells % (No.) Maternal Age by Female and Polar Bodies Analyzed Years (Range) (Cycles) MII 1PB 2PB MII 1PB 2PB

(Dailey, Dale, Cohen, & Munne, 1996) 26-34 DNA 13, 18, 21, X 383 188 CNA Unfertilized 2.9 (2/67) DNA DNA 35-39 13.2 (9/68) >40 30 (10/33) (Verlinsky et al., 1996) >35 135 13, 18, 21, X 648 251 128 Normally 32.1 30.2 19.1 Fertilized (208/648) (157/251) (75/128) (Verlinsky et al., 1997) >35 395 (398) 13, 18, 21 3651 CNA Normally 43.1 (1271/3651) DNA Fertilized (Verlinsky et al., 1998) >35 363 (538) 13, 16, 18, 21, 22 2742 CNA Normally 40 (1102/2742) DNA Fertilized (Martini et al., 2000) 32 (26-39) 18 (18) 1, 7, 13, 18, 21, X 57 CNA CNA Unfertilized 43.5 (25/57) DNA DNA (Mahmood, Brierley, Faed, Mills, & 33 72 1, 9, 13, 16, 18, 21, X 127 57 CNA Unfertilized and (6/127) (3/57) DNA Delhanty, 2000) in vitro matured (Munne et al., 2000) 35.2 (29-39) 6 13, 16, 18, 21, 22 87 CNA Normally 33.3 (29/87) DNA Fertilized (Verlinsky et al., 2001) (35-45) 585 (917) 13, 16, 18, 21, 22 CNA 4016 3895 Normally 45.2 36.1 29.3 Fertilized (2077/4599) (1449/4016) (1143/3895) (Honda et al., 2002) 31.7 (23-45) 84 18, 21, X 183 93 CNA Unfertilized 3 (5/183)1 6.5 (6/93) CNA (Benkhalifa, Kahraman, Caserta, (22043) 507 13, 16, 18, 21, 22, X 711 CNA CNA Unfertilized 20.5 DNA DNA Domez, & Qumsiyeh, 2003) (183/711) (Cupisti et al., 2003) 32.5 (22-44) 124 1, 9, 12, 13, 16, 18, 21, X 230 88 CNA Unfertilized 4.3 (10/230) 3.4 (3/88) DNA (Pujol et al., 2003) 33.7 (18-45) 60 1, 13, 15, 16, 17, 18, 21, 89 73 CNA In vitro matured 68.5 (61/89) 54.8 (40/73) DNA 22, X (Magli et al., 2004) 38.4 48 (51) 13, 15, 16, 18, 21, 22, X, Y 346 CNA Normally 53 (183/346) DNA Fertilized (Gutierrez-Mateo et al., 2005) 32.9 (6-42) 34 mFISH 16 14 CNA In vitro matured 25 (4/16) DNA DNA (Montag et al., 2005) 39.1 (75) 13, 16, 18, 21, 22, X 477 CNA Normally 53.7 DNA 38.1 Fertilized (256/477) (Magli et al., 2006) 37.8 75 13,16,18,21, 22 564 CNA Normally 58.6 54 (197/366) DNA (87) Fertilized (In vitro (309/527) 70 (112/161) matured) (Vlaisavljevic, Krizancic Bombek, 32.9 173 13,16,18,21, 22 131 CNA In vitro matured 45 (59/131)3 DNA DNA Vokac, Kovacic, & Cizek-Sajko, 2007) (22-43) 81 48.1 (39/81)3 (Gianaroli et al., 2010) 38.2 544 13,16,18,21, 22 4163 CNA Normally 49.3 DNA (38-45) (706) Fertilized (1885/3816) (Kuliev et al., 2011)1 38.8 2830 13,16,18,21, 22 20986 Normally 46.8 30.4 39.8 (35-45) (3953) Fertilized (9812/20986) (2983/9812) (3908/9812) (Magli et al., 2012) 33.3 51 13,16,18,21, 22 259 Normally 12.4 25.1 12.4 (38-45) (51) Fertilized (32/259) (65/259) (45/259) (Hammoud et al., 2012) >36 28 13,16,18,21, 22 156 CNA Normally 35.8 35.3 (55/156) DNA <36 17 101 Fertilized (112/313) 31.7 (32/101) >36 15 56 44.6 (25/56) DNA refers to “data not available” which were not mentioned by the author in the study, and CAN refers to “cell not analyzed”. 1 The study is combined with the data of (Kuliev, Cieslak, Ilkevitch, & Verlinsky, 2003). 2 The rates of two different groups of maternal age. 3 The rates of two different groups of in vitro matured oocytes.

5 Table 1.1.-C: CGH Analysis Studies Average No. of Female No. of Analyzed oocytes Most Chromosomes Overall Rate of Aneuploid Overall Aneuploidy Rate Maternal Age (Cycles) and first polar bodies Involved in Aneuploidy Cells % (No.) of oocytes in % (No.) Years (Range) MII-1PBs MII 1PB (Gutierrez-Mateo, Wells, et al., 2004) 33.2 (21-41) 21 25 17, 19, 22 48 (12/25) 48 (12/25) 48 (12/25)1 (Gutierrez-Mateo, Benet, et al., 2004) 34.9 (23-42) 46 42 1, 4, 13, 16, 22 88.1 (37/42) 88.1 (37/42) 88.1 (37/42) (Magli et al., 2004) 38.4 110 (113) 260 DNA 53 (183/260) 47.9 (114/238) 53 (183/260) (Fragouli, Wells, Whalley, et al., 2006) 32.2 15 15 13, 16, 21, 22 66.7 (10/15) 62.5 (10/16) 66.7 (10/15) (Fragouli, Wells, Thornhill, et al., 2006) 32.5 (18-42) 15 100 8, 13, 20, 21, X 22 (22/100) 22 (22/100) 22 (22/100) (Landwehr, Montag, van der Ven, & Weber, 2008) 38 (33-44) 16 32 6, 7, 8, 11, 19 75 (24/32) 75 (24/32) 75 (24/32) (Fragouli et al., 2009) 22 13 121 DNA 3.3 (4/121) 3.3 (4/121) 3.3 (4/121) (Fragouli, Alfarawati, Goodall, et al., 2011) 40.8 (34-47) 70 308 DNA 70 (219/308) 40 (124/308) 70 (219/308) DNA refers to “data not available” which were not mentioned by the author in the study. All studies analyzed a mixture of unfertilized oocytes and in vitro matured oocytes. 1 The overall rates of aneuploidy showed a statistical difference in relation to maternal age (two groups of women were 37 years old or younger, and women were older that 37 years).

Table 1.1.-D: aCGH and SNP Analysis Studies Maternal No. of No. of Successfully No. of Meiotic Errors (%) Most No. of Aneuploid PBs Overall Aneuploidy Age Mean or Female Analyzed PBs (%) Chromosomes (%) Rate of oocytes in % Average in (Cycles) Involved in (No.) Years Aneuploidy 1PBs 2PBs MI MII MI&MII 1PBs 2PBs (Fishel et al., 2011) 41.0 (38-43) 134 (150) 861 CNA 689 DNA DNA 15, 16, 21, 22 580 (67.4) CNA 67 (580/689) (Gabriel et al., 2011) (29-50) 25 164 (97) CNA 256 DNA DNA 15, 21, 22 86 (52.4) CNA 52 (86/164) (Geraedts et al., 2011) 40.0 41 212 (94) 207 (92) DNA DNA DNA DNA 122 (58) 142 (69) 72 (140/195) (Handyside et al., 2012) 40.0 (33-44) 34 105 105 77 (34) 102 (45) 50 (14) 16, 22, 21, 19 DNA DNA 76 (80/105) (Christopikou et al., 2013) 39.0 20 92 (100) 54 (93) 30 (30) 48 (48) 22 (22) DNA DNA DNA 72 (39/54) (Capalbo et al., 2013) 43.0 (40-45) 9 (9) 45 (92) 41 (84) 9 (12) 55 (71) 14 (18) 22, 15, 16, 17 DNA DNA 52 (23/42) (Salvaggio, Forman, Garnsey, Treff, & 35.3 (24-42) 96 440 442 DNA DNA DNA DNA 110 (25) 122 (27.6) 41 (179/433) Scott, 2014) DNA refers to “data not available” which were not mentioned by the author in the study, and CAN refers to “cell not analyzed”.

6 The maternally related chromosome abnormalities may also occur prior to meiotic divisions as pre-meiotic mitotic errors. It has been proposed that these errors arise as a result of mitotic errors occurring during germ cell proliferation in which chromosomally abnormal cells may ultimately become gametes that carry the same chromosomal error. It has also been hypothesized that pre-meiotic mitotic errors may arise during preimplantation embryonic development in which chromosomally abnormal cells may eventually become precursors to germ cells; both aforementioned hypotheses are illustrated in Figure 1.1 and have been reviewed recently (Biesecker & Spinner, 2013; Delhanty, 2011; Taylor et al., 2014). These pre-meiotic mitotic errors can be defined as germline/gonadal mosaicism. The majority of the studies on gonadal mosaicism to date have been limited to only young women with successive conceptions with trisomy (Bruyere, Rupps, Kuchinka, Friedman, & Robinson, 2000; Frias, Ramos, Molina, del Castillo, & Mayen, 2002; Harris, Begleiter, Chamberlin, Hankins, & Magenis, 1982;

Kovaleva, 2010; Pangalos et al., 1992; Sachs, Jahoda, Los, Pijpers, & Wladimiroff, 1990; Tseng,

Chuang, Lee, & Ko, 1994; Uchida & Freeman, 1985), giving rise relatively low rate of gonadal mosaicism of about 2 to 10% (Papavassiliou, Charalsawadi, Rafferty, & Jackson-Cook, 2015).

To date, the contribution of pre-meiotic mitotic errors to the overall rate of chromosome abnormalities in the general population, the clinical relevance to human reproduction and fertility, and the association with maternal age remain unknown. Some studies have been conducted to investigate the prevalence and cytogenesis of pre-meiotic mitotic errors in human foetal ovarian cells obtained from women with normal karyotypes; all these studies are listed in

Table 1.2-A. One of these studies has proposed a model called the Oocyte Mosaicism Selection

Model (OMSM) which suggests that pre-meiotic mitotic errors are maternal-age dependent

(Hulten, Patel, Jonasson, & Iwarsson, 2010). In this study, it was hypothesized that from the

7 relatively high rate of trisomic germ cells in the foetal ovaries with normal karyotype, many may be able to initiate the prophase stage and progress to the dictyate stage. At the time of adulthood, selective selection against these oocytes for ovulation may occur, resulting in an increased pool of chromosomally abnormal oocytes within the antral follicle count among women of advanced age. Only two research groups re-examined the first half of the proposed model, but both failed to replicate Hulten’s model (Morris et al., 2012; Rowsey et al., 2013).

Only one study to date has investigated the incidence of pre-meiotic mitotic errors in human mature oocytes with the corresponding first polar bodies (Obradors et al., 2010) (Table

1.1-B). Another study has been conducted by the same research group analyzing the incidence of pre-meiotic mitotic errors in human immature oocytes retrieved from women who underwent

ART treatment cycles (Table 1.1-C) (Daina et al., 2014). Inconsistent results were reported in the overall rates of pre-meiotic mitotic errors in immature and mature oocytes in both studies.

Observations from limited studies show a great discrepancy in the overall rate of pre- meiotic mitotic errors in human mature oocytes with the corresponding first polar bodies derived from women who underwent ART treatment cycles. This is largely because these studies were not intended to investigate this type of chromosomal error (Table 1.2-B). Most of these studies were designed to develop and standardize a larger panel of fluorescence in-situ hybridization

(FISH) probes or comparative genomic hybridization (CGH) techniques for chromosomal abnormality screening in mature oocytes and polar bodies. A high rate of experimental artifacts, as well as the small sample size, mixed population of oocytes (unfertilized oocytes and in vitro matured oocytes which were not subjected to sperms), and the diverse population of female subjects (donors or patients with fertility problems), are all contributing factors to the incidence of pre-meiotic mitotic errors.

8

Figure 1.1: The proposed hypotheses for the origin of pre-meiotic mitotic errors in human oocytes. The first proposed hypothesis states that the chromosomally abnormal immature oocytes may arise as a result of mitotic errors during germ cell proliferation within the foetal ovaries. Germ cells carrying these pre-meiotic mitotic errors may ultimately become chromosomally abnormal gametes within the adult ovaries. At the time of ovulation, one of these chromosomally abnormal immature oocytes may undergo the first meiotic division, giving rise to a chromosomally abnormal zygote. According to the second hypothesis, chromosomally abnormal immature oocytes arise as a result of mitotic errors during the preimplantation stage of embryonic development, either at cleavage or blastocyst formation. The chromosomally abnormal cells within the mosaic embryos may ultimately become an adult female with either gonosomal or gonadal mosaicism whose ovaries consist of a pool of chromosomally abnormal germ cells, which may ultimately become chromosomally abnormal gametes (parental generation). At the time of ovulation, one of these chromosomally abnormal immature oocytes may undergo the first meiotic division, giving rise to a chromosomally abnormal zygote (second generation). The rightmost panel illustrates a chromosomally abnormal immature oocyte undergoing the first meiotic division to give rise to a chromosomally normal mature oocyte and ultimately, a chromosomally normal zygote.

Part of the figure was adopted from (Biesecker & Spinner, 2013).

9 Table 1.2: Summary of the overall rates of pre-meiotic mitotic errors that have been investigated in human foetal ovarian cells and adult oocytes using different techniques for analysis. Table 1.2-A: The overall rate or pre-meiotic mitotic errors in foetal ovarian cells. Studies No. Ovarian Samples Gestational week No. of Analyzed Cells Overall Rate of Pre-Meiotic Overall Rate of Chromosome Technique Mitotic Errors Rate (range) Normality (range) (Hulten et al., 2008) 8 CN 14-22 w 2126 PM cells 0.8% (17/2126) Chr.21 (2,109/2126) Chr.21 FISH dual probes for Chr.21 3653 M cells 0.66% (24/3653) Chr.21 (3,629/3653) Chr.21 (Morris et al., 2012) 8 CN 10-14 w 9085 cells 0% (0/9085) Chr.163 100% (9085/9085) Chr.16 FISH for Chr.16, 21, and 22 8365 cells 0.02% (2/8365) Chr.213 99.9% (8363/8365) Chr.21 8645 cells 0.01% (1/8645) Chr.223 99.9% (8644/8645) Chr.22 (Rowsey et al., 2013) 7 CN 16-23 w 1,034 cells 0%5 66.3% (686/1,034) Chr.16 FISH for Chr.13, 16, and 21 1,206 cells 62.3% (758/1,206) Chr.13, 21 IF (confirmation PCA) (Hulten et al., 2014) 12 CN 9-11 w 27,115 cells 0.066% (18/27,115) 99.73% (27,042/27,115) FISH dual probes for Chr.21

Table 1.2-B: The overall rate or pre-meiotic mitotic errors in human mature oocytes with the corresponding polar bodies. Studies No. of No. of Patients Type of Samples Overall rate of Meiotic Overall Rate of Pre-meiotic Technique Analyzed Cells (Mean, range age yrs.) Errors (No. of Abnormal over Errors (No. of Abnormal over Total oocytes) Total oocytes) (Obradors et al., 2010) 84 MII-1PBs 53 IVF donors (26.1) Mixed of in vitro matured 17.8% (15/84) 15.5% (13/84) CGH and unfertilized oocytes (Fragouli, Wells, 15 MII and 15 IVF patients (32.2) Mixed of in vitro matured 55.5% (5/9) 11.1% (1/9) CGH Whalley, et al., 2006) 161PBs and unfertilized oocytes (9 MII-1PBs) (Fragouli, Wells, 100 MII-1PBs 46 IVF patients Mixed of in vitro matured 22% (22/100) 1% (1/100) CGH Thornhill, et al., 2006) (32.5) and unfertilized oocytes (Gutierrez-Mateo, Benet, 42 MII-1PBs 46 IVF patients Mixed of in vitro matured 88.1% (37/42) 9.5% (4/42) CGH-FISH for chr.13, 16, 18, et al., 2004) (34.9) and unfertilized oocytes 21 and 22 (Gutierrez-Mateo, Wells, 25 MI-1PBs 21 IVF patients Mixed of in vitro matured 48% (12/25) 24% (6/25) CGH et al., 2004) (33.2) and unfertilized oocytes (Cupisti et al., 2003) 230 MII and 88 124 IVF patients Mixed of in vitro matured 4.3% (10/230) and 3.4% (3/88) 1.3% (3/230) FISH for chr.1, 9, 12, 13, 16, 1PBs (32.5, 22-44) and unfertilized oocytes 18, 21and X (Pujol et al., 2003) 46 MII-1PBs 60 IVF patients In vitro matured, 41.3% (19/46) 23.9% (11/46) FISH for chr.1, 13, 15, 16, 17, (33.7, 18-45) unfertilized oocytes 18, 21, 22 and X (Mahmood et al., 2000) 127 MII and 57 72 IVF patients Mixed of in vitro matured 5.5% (7/127) and 5.3% (3/57) 3 MII-1PBs FISH for chr.1, 9, 13, 16, 18, 1PBs (33) and unfertilized oocytes 21 and X

(Cozzi et al., 1999) 4 0PN-1PBs 1 IVF patient (37) Unfertilized oocytes 100% (4/4) 75% (3/4) FISH for chr.21

Table 1.2-C: The overall rate or pre-meiotic mitotic errors in human immature oocytes. Studies No. of Analyzed Cells No. of Patients Type of Samples Overall Rate of Pre-meiotic Technique (Mean, range age yrs.) Errors (No. of Abnormal over Total oocytes) (Daina et al., 2014) 157 GV/MI 32 IVF patients (25-45) and Immature oocytes (GV/MI) 15.3% (24/157) CGH 24 IVF donors (18-33) (Garcia-Cruz, 40 MI 140 IVF donors Immature oocytes (MI) 25% (10/40) M-FISH and IF Casanovas, et al., 2010) (26.6, 18-35)

10 1.1.2. The Rationale of the Study

Most of the studies investigated the incidence of germline mosaicism were limited to specific group of young women with trisomic conceptions (Papavassiliou et al., 2015). Only a handful of studies have been conducted to investigate the prevalence and cytogenesis of germline mosaicism in human foetal ovarian cells due largely in part to the difficulty obtaining these samples from fetuses with normal karyotypes (Table 1.2). The Oocyte Mosaicism Selection

Model (OMSM) suggests that pre-meiotic mitotic errors are maternal-age dependent (Hulten et al., 2010). The relatively high rate of trisomic germ cells in foetal ovaries may be able to initiate the prophase stage and progress to the dictyate stage. During reproductive life, selective selection against these oocytes for ovulation may result in the increased number of chromosomally abnormal oocytes within the antral follicle count amongst women of advanced age compared to younger women. Only two research groups re-examined the first half of the proposed model

(Morris et al., 2012; Rowsey et al., 2013), and failed to replicate Hulten’s group findings.

One study has analyzed the incidence of pre-meiotic mitotic errors in human mature oocytes with the corresponding first polar bodies (Obradors et al., 2010). Another study has been conducted by the same research group to investigate the incidence of these errors in human immature oocytes retrieved from women who underwent ART treatment cycles (Daina et al.,

2014). Although both studies have been conducted by the same investigators, there were inconsistent results in relation to the incidence of women who produced chromosomally abnormal oocytes due to pre-meiotic mitotic errors. Other limited studies reported observations on pre-meiotic mitotic errors in human mature oocytes with the corresponding first polar bodies derived from women who underwent ART treatment cycles (Table 1.2-B). However, these

11 observations showed high discrepancy in the overall rate of pre-meiotic mitotic errors, as these studies were not intended to investigate this type of chromosomal errors.

Despite all these limited studies, the contribution of pre-meiotic mitotic errors to the overall rate of chromosome abnormalities in the general population, the clinical relevance to human reproduction and fertility, and the association with the maternal age remain unknown which are intriguing scientific questions that we feel is important to explore.

1.1.3. Objectives of the Project

Therefore, the main objective of our project was to investigate the incidence of pre-meiotic mitotic errors in human immature and mature oocytes derived from women who underwent controlled ovarian stimulation (COH) treatment cycles in relation to maternal age. FISH analysis was performed for those chromosomes most frequently involved in pregnancy loss. In order to fulfill this aim, the following specific objectives were successfully achieved.

1.1.3.1. Specific Objectives

The first objective was aimed to optimize the FISH technique for the chromosomes (13, 15,

16, 18, 21, 22 and X) using interphase nuclei obtained from human lymphocytes (as a negative control) and human spare, chromosomally abnormal preimplantation embryos (as a positive control).

The second objective was aimed to analyze the chromosomal complements in human immature and mature oocytes using the standardized FISH technique.

The third objective was intended to validate FISH signals for all chromosomes analyzed in immature oocytes at different maturation stages, as well as in mature oocytes (as an internal control) by plotting histogram distributions of FISH signals per tested chromosome of each

12 oocyte group in relation to maternal age. Chai-squared test was used to determine the statistical

difference between the histograms between both maternal age groups.

The fourth objective was designed to estimate the overall rate of pre-meiotic mitotic errors

in relation to maternal age by investigating the incidence of chromosomally normal immature

oocytes per maturation stage group. Concomitantly, the incidence of chromosomally normal

mature oocytes and unfertilized oocyte groups was performed for comparison.

The fifth objective estimated the overall rate of pre-meiotic mitotic errors in association to

maternal age by re-analyzing the incidence of chromosomally normal immature and mature

oocytes from the same cohort when each patient donated at least one immature oocyte at any

maturation stage and one mature oocyte (sibling oocytes) for elimination the patient-to-patient

variation bias.

The sixth objective investigated the incidence of pre-meiotic mitotic errors in mature

oocytes, including unfertilized oocytes with the corresponding first polar bodies in relation to

maternal age. Further analysis of the incidence of first meiotic errors in mature and unfertilized

oocytes with the corresponding first polar bodies was accomplished for comparison.

1.1.4. Significance of the Project

To our knowledge, the present study is the first to analyze the chromosomal complement in

such a considerable sample size using FISH analysis. Our study analyzed human immature

oocytes at different maturation stages, including mature and unfertilized oocytes with their

corresponding first polar bodies. This work reported for the first time the prevalence of pre-

meiotic mitotic errors in sibling oocytes. Observations on different patterns of chromosomal

segregation and leading mechanisms to pre-meiotic mitotic errors in mature oocytes were

reported in this study.

13 1.2. LITERATURE REVIEW

The literature to date, including substantive findings and methodological contributions to chromosomal abnormalities in human oocytes in association to maternal age, is going to be presented in the following section.

1.2.1. Characteristics of Chromosome Abnormalities in Humans

Discovery of Chromosome Abnormalities in Humans

The human karyotype in human gametes was first determined in 1956 (C. E. Ford &

Hamerton, 1956), in which the normal diploid (2N) human cell was identified to contain 46 chromosomes (23 homologous pairs). Three years later, the chromosomal complement of three clinical cases of Down syndrome was first described (Lejeune, Turpin, & Gautier, 1959) with karyotypical analysis showing an extra copy of chromosome 21 (trisomy 21) in the somatic cells.

Soon after, the chromosomal aetiology of this condition was confirmed (P. A. Jacobs, Baikie,

Court Brown, & Strong, 1959). Further discoveries of related chromosomal aetiologies of some other genetic conditions were reported soon after, including Klinefelter syndrome (47, XXY) (P.

A. Jacobs & Strong, 1959), Turner syndrome (45, XO) (C. E. Ford, Jones, Polani, De Almeida,

& Briggs, 1959), Triple-X syndrome (47, XXX) (P. A. Jacobs, Baikie, Brown, et al., 1959),

Edward syndrome (Trisomy 18) (Edwards, Harnden, Cameron, Crosse, & Wolff, 1960), Patau syndrome (Trisomy 13) (Smith, Patau, Therman, & Inhorn, 1960), and XYY syndrome (47,

XYY) (Muldal, Ockey, Thompson, & White, 1962).

All these discoveries used uniformly stained metaphase chromosomes (under light microscope) to visualize the abnormalities. The chromosomes were classified into seven groups,

A to G, based on their length and centromere position (C. E. Ford & Hamerton, 1956; Tjio,

1978). Since the initial discoveries, advances in chromosomal staining techniques have

14 facilitated the identification of the individual chromosome and any alterations in their number and/or structure (Caspersson et al., 1968; Holmquist, 1992).

Characteristics of Chromosome Abnormalities in Humans

There are two main types of chromosomal abnormalities in humans: numerical and structural abnormalities.

Numerical chromosome abnormalities are the most common type in which one or more of chromosomes are numerically imbalanced. There are different types of numerical chromosome abnormalities, such as aneuploidy, haploidy, and polyploidy. Aneuploidy is the most common type of numerical chromosome abnormalities in which only one or two of the chromosomes are lost or gained; cells with missing chromosomes are monosomic, while those with an extra chromosome are trisomic. For example, Down syndrome (Trisomy 21) is a well-known condition caused by an additional copy of chromosome 21. Haploidy and polyploidy are less frequent compared to aneuploidy. Cells with one copy of each chromosome are referred to as haploid (1N), while cells with an extra copy of each chromosome are polyploid. These polyploid cells can be triploid (3N) and tetraploid (4N) when these cells contain three and four copies of each chromosome, respectively (Griffiths et al., 2000).

In all the previously mentioned types of numerical chromosome abnormalities, cells are all uniformly aneuploid, haploid, or polyploid. However, cells can be defined as mosaic when diploid cells co-exist with any non-diploid cells, including various combinations of aneuploid, haploid, or polyploid cell populations within a single preimplantation embryo or individual. Cells also can be chaotic when multi-chromosomal losses or gains of all cells exist (Bielanska, Jin,

Bernier, Tan, & Ao, 2005; Bielanska, Tan, & Ao, 2002a, 2002b, 2003); this occurrence is a sub- category of mosaicism. All these conditions were observed for the first time in healthy

15 individuals in the mid-20th century (Cotterman, 1956). Subsequently, the frequency of mosaic cases was estimated in viable pregnancies (Hsu & Perlis, 1984), spontaneous abortions (T.

Hassold et al., 1980), foetal and/or placental tissues (Crane & Cheung, 1988), and human preimplantation embryos (Plachot et al., 1987).

Structural chromosomal abnormalities are less common as compared to numerical anomalies. This can involve a chromosomal breakage with or without partial loss of one or two of the chromosomes involved. There are different types of structural chromosome abnormalities including deletions, duplications, inversions, insertions, and translocations. Translocations are the most common type of structural chromosome abnormality and are subcategorized into reciprocal and Robertsonian translocations. In reciprocal translocation, an exchange of chromosomal material occurs between two chromosomes resulting in either balanced reciprocal translocation with no chromosomal material loss, or unbalanced reciprocal translocation with chromosomal material loss. Whereas, in Robertsonian translocation, extra chromosomal material is lost or gained when two of the acrocentric chromosomes (13, 14, 15, 21, or 22) are rearranged

(Griffiths et al., 2000).

The ability to identify human numerical and structural chromosomal abnormalities, has allowed for research to investigate the incidence and the clinical relevance of these abnormalities in human reproduction.

1.2.2. Significance and Incidence of Chromosome Abnormalities in Human Reproduction

The Overall Rate of Chromosome Abnormalities in Human Reproduction

The negative impact of numerical chromosome abnormalities on human reproduction has been well documented for over 50 years (Hamerton et al., 1975). Today, these anomalies are known to be the leading cause of very early embryo loss, spontaneous abortion, stillbirths, and

16 birth defects (J. H. Ford et al., 1996; T. J. Hassold, 1986; T. J. Hassold & Jacobs, 1984; Plachot et al., 1986, 1987; Spandorfer et al., 2004). It has been estimated that around 10% of all clinically recognized pregnancies are have chromosomally abnormal products of conception. The incidence of these abnormalities varies enormously during the developmental time point, where approximately 0.3% of live births (with congenital malformations), and 4% of stillbirths are due to chromosome abnormalities. This estimated rate of chromosome abnormalities increases to around35% of clinically recognized spontaneous abortions (T. Hassold & Hunt, 2001).

The overall Rate of Chromosome Abnormalities in Live Births

Only autosomal trisomies for chromosomes 21, 18, 13, and sex chromosomes are compatible with life, and may result in live births with congenital abnormalities. Autosomal trisomies for the remaining chromosomes, as well as most monosomies (with the exception of

45, XO) are lethal. Amongst the most common, non-lethal trisomies, the prevalence rate of

Trisomy 21 is the highest, at around 0.13% of newborns, while the prevalence rate of trisomy 13 is the lowest at about 0.005% of newborns. The prevalence rate of monosomy of the X chromosome (Turner syndrome 45, XO) is about 0.01% of newborns (Hamerton et al., 1975; T.

J. Hassold, 1986; T. J. Hassold & Jacobs, 1984). Trisomy 21 and sex chromosome abnormalities are the most compatible with postnatal survival, while Trisomy 18 and Trisomy 13 are the least.

The clinical manifestations of trisomy 21 include growth and mental retardation, and congenital malformations (Hamerton et al., 1975); whereas, infertility, various degrees of mental impairment, and dysmorphic characteristics are associated with sex chromosome abnormalities

(Polani, 1969a). Trisomy 18 and 13 are associated with more severe anatomical defects that are lethal shortly after birth (Carter, Pearn, Bell, Martin, & Anderson, 1985; Moerman, Fryns, van der Steen, Kleczkowska, & Lauweryns, 1988).

17 The overall Rate of Chromosome Abnormalities in Stillbirths

In addition to the anomalies involved in live births, autosomal trisomies for chromosomes

21, 18, 13, and sex chromosomes may result in foetal death between the 20 week of gestation and birth. The prevalence rates in stillbirths for trisomies of chromosomes 21, 18, 13, and sex chromosomes are 1.3%, 1.1%, 0.3%, and 0.1% respectively. Autosomal trisomies for the rest of the chromosomes, as well as majority of the monosomies are lethal, except for monosomy of the

X chromosome (45, XO) with a prevalence of about 0.1% of pregnancies ending in stillbirths.

Triploidy, on the other hand, is reported in stillbirth to have a prevalence rate of about 0.2% (T.

J. Hassold, 1986; T. J. Hassold & Jacobs, 1984).

The overall Rate of Chromosome Abnormalities in Spontaneous Abortions

In contrast to those anomalies involved in live births (with congenital defects) and stillbirths (pregnancy loss after 20 weeks gestation), chromosomal abnormalities are more prevalent among clinically recognized spontaneous abortions (foetal death between 6 to 20 weeks of gestation) and report a frequency rate of about 50%. The most common types of these chromosome abnormalities are monosomies for chromosome X, and trisomies for chromosome

16 with estimated rates of about 8.7% and 7.5%, respectively. Also, triploidy and tetraploidy lead to early pregnancy loss with frequency rates of 6.4% and 2.4%, respectively. These rates are followed by trisomies for chromosome 22 (2.7%), and for chromosomes 2, 13, 14, 15, and 21

(1% to 2.3%). Trisomies for chromosomes 3, 5, 6, 11, 12, 17, and 19 are less frequent, while monosomies for all autosomal chromosomes are the least frequent. Trisomy for chromosome 1 has not been reported to date (J. H. Ford et al., 1996; T. J. Hassold, 1986; T. J. Hassold & Jacobs,

1984).

18 The overall Rate of Chromosome Abnormalities in Pre-clinical Abortions, Pre-implantation

Embryos, and Gametes

It is believed that the overall rate of chromosome abnormality remains underestimated where only clinically recognized pregnancies have been included. However, cytogenetic analyses of pre-clinical abortions, which occurred prior to 6 weeks of gestation, were mostly excluded (T.

Hassold & Hunt, 2001). Advances in ART have a significant role in assessing the rate of chromosome abnormalities at very early stage of pregnancies. The improvements in preimplantation genetic diagnosis (PGD) for chromosome abnormality screening have a substantial role in investigating the overall rates of chromosome abnormalities in human gametes and pre-implantation embryos (Brown & Harper, 2012; Diamond, Willman, Chenette, & Cedars,

2012).

The overall rate of chromosome abnormalities in spontaneous abortions of early pregnancy loss

Karyotype analysis has revealed that around 74% of ART produced conceptions are lost prior to 6 weeks of gestation due to chromosome abnormality (Spandorfer et al., 2004). The most common types of these chromosome abnormalities are trisomies for chromosome 21, 16, 22, 15, and 18 with frequency rates of about 23.9%, 13%, 10.9%, 10.9% and 8.7%, respectively. These rates are followed in frequency by monosomy X (3.5%), trisomy XXY (2.4%), and triploidy

(1.8%).

The overall rate of chromosome abnormalities in human preimplantation embryos

The frequency of chromosomally abnormal human preimplantation embryos is around

75% for cleavage stage embryos, and 50% for blastocysts, with a wide range of reported results

(Fragouli et al., 2013; Fragouli & Wells, 2011; Mantikou, Wong, Repping, & Mastenbroek,

2012). A recent systemic review and meta-analysis of 36 studies investigated the chromosomal

19 complements of human preimplantation embryos using different methodologies (van Echten-

Arends et al., 2011). The review considered three main factors for the meta-analysis: the number of chromosomes analyzed, the method applied for analysis, and the developmental stage studied of the embryos (cleavage stage or blastocyst stage). The authors found that 73% of all 815 monospermic, IVF-generated embryos are mosaic, of which 59% and 15% of the total were diploid-aneuploid mosaic and aneuploid mosaic embryos, respectively. Surprisingly, only 39 out of 815 embryos (5%) were chromosomally abnormal, including aneuploid, haploid, and polyploid embryos. All chromosomes have been analyzed in preimplantation embryos at both the cleavage and blastocyst stages, with some chromosomes having a higher prevalence of monosomies or trisomies for chromosomes 13, 16, 21, and 22 (Fragouli et al., 2013).

The overall rate of chromosome abnormalities in human gametes

On average, it has been estimated that about half of normally fertilized oocytes are chromosomally abnormal (Kuliev et al., 2011), and this directly correlates with advanced maternal age. The contribution of errors occurring in either meiosis I (MI) or meiosis II (MII) (to the overall rate of chromosome abnormality per oocyte) is found to be relatively equal (Kuliev et al., 2011).

Studies spanning two decades have shown a wide range in the overall rate of chromosome abnormalities (meiotic errors) in human oocytes, as listed in Table 1.1. There are several factors that may contribute to the heterogeneity in reported meiotic error rates in human oocytes in the literature. Maternal age, the chromosome analyzed, the number of chromosomes screened, the methodology applied, the type of oocytes studied, the hormonal stimulation regime applied, and the type of fertility issue involved are all factors have been shown to affect the overall rate of chromosome abnormalities in human oocytes (Table 1.1) (Fragouli, Wells, et al., 2011; B.

20 Rosenbusch, Schneider, & Michelmann, 2008). A systemic review and meta-analysis on the chromosomal complements of human mature oocytes is required in order to clarify further this heterogeneity.

Meiotic errors are chromosome-specific. For example, almost all cases of oocytes with trisomy 16 are MI errors, while cases with trisomy 18 are mostly MII errors. Oocytes with trisomy 15, 21, and 22 are more likely due to MI errors with a rate three times higher than MII errors (T. Hassold et al., 2007).

Recent studies are trying to increase the number of chromosomes being screened using

FISH analysis in PBs (Cupisti et al., 2003; Gutierrez-Mateo et al., 2005; Pujol et al., 2003), or are using more developed methodologies such as CGH (Fragouli, Wells, Whalley, et al., 2006), aCGH (Gabriel et al., 2011; Handyside et al., 2012; Jaroudi & Wells, 2013), and SNP array

(Scott et al., 2012) for comprehensive chromosome screening (CCS) in PBs (Pellestor et al.,

2005; Treff & Scott, 2012). Screening chromosome abnormalities in polar bodies is challenging, compared to analyzing single biopsied blastomeres from cleavage stage embryos of trophectoderm cells (TE) which have been well standardized (Harper & Harton, 2010). This prompted The European Society of Human Reproduction and Embryology (ESHRE) takes the initiative to develop and standardize chromosome abnormality screening in PBs using aCGH, called ESHRE PGS TASK FORCE (Geraedts et al., 2010). This task is divided into two main parts; the first, which has already been conducted, is a pilot study to assess the efficacy of chromosome abnormality screening in PBs using aCGH (Gabriel et al., 2011). The second part, which is ongoing, is a multicenter randomized control trial to determine whether chromosome abnormality screening in PBs using aCGH increases the delivery rate in women of advanced

21 maternal age. The outcome from this trial will answer whether this approach has a clinically significant advantage over the cleavage or blastocyst biopsy.

The type and source of human oocytes analyzed have a possible impact on the overall rate of meiotic errors, although a definitive association between oocyte characteristics and meiotic error rates remain undefined. Some studies have revealed that the type of oocyte studied may be associated with an increased rate of meiotic errors (Schmutzler et al., 2014), but not the morphology of polar bodies (Treff et al., 2012). Unfertilized oocytes (which failed to fertilize in vitro after insemination or ICSI) are found to be more associated with chromosome abnormalities (B. Rosenbusch et al., 2008) as compared to normally fertilized oocytes. Other research groups have found that unfertilized oocytes are significantly associated with chromosome fragmentation, premature chromosome condensation, and cytoskeletal defects; these findings were also analyzed in correlation with the subject’s age and infertility history

(Zhivkova, Delimitreva, Toncheva, & Vatev, 2007). Another study observed a higher frequency of complete or partial premature chromatid separation in unfertilized oocytes (B. E. Rosenbusch

& Schneider, 2006). The rate of meiotic errors are found to be comparable between human oocytes matured in vivo or in vitro, either as leftover from stimulated cycles or generated from in vitro maturation (IVM) treatment cycles (Coticchio et al., 2013; Vlaisavljevic et al., 2007; Yakut,

Karkucak, Sher, & Keskintepe, 2012). However, the prolonged time of in vitro maturation has been found to be significantly associated with spindle disorganization and chromosome misalignment (Lei, Guo, Liu, Tan, & Li, 2014; Y. Li et al., 2006; Yu et al., 2011), but not with an altered rate of meiotic errors (Vieira et al., 2011; Vlaisavljevic et al., 2007).

Approximately 70 to 80% of all monosomy XO cases are of paternal of origin; while, around 50% and 100% of trisomy XXY and trisomy XYY are paternally contributed,

22 respectively (P. Jacobs et al., 1997; Martin et al., 1983; Martin & Rademaker, 1999).

Chromosomal analysis has been performed on over 10 000 sperm nuclei collected from men from the general population ranging in age from 10 to 80 years (Templado et al., 2013). From this analysis, disomy of the sex chromosome is the most frequent, followed by disomy of chromosome 21 with a prevalence of about 0.27% and 0.17%, respectively.

Overall, the combined data from in utero conceived clinically recognized pregnancies, and assisted conceived pregnancies, indicate a significantly high level of chromosome abnormalities

- at least 5% and possibly as high as 25% of all clinically recognized pregnancies (T. Hassold &

Hunt, 2001). In addition, these results reveal that numerical chromosome abnormalities are not only the leading genetic cause for spontaneous abortion, stillbirths and live births with congenital defects, but also the underlying cause of very early pregnancy loss, implantation failure and embryos wastage (Fragouli, Alfarawati, Spath, & Wells, 2014; Pehlivan et al., 2003; Rubio et al.,

2013).

The overall rate of structural chromosome abnormalities

The incidence of structural chromosome abnormalities is much less as compared to numerical anomalies. Less than 1% of live birth defects are due to different types of imbalanced structural rearrangements; however, about 3% are phenotypically normal individuals who are chromosomally balanced (T. J. Hassold, 1986; T. J. Hassold & Jacobs, 1984; P. A. Jacobs, 1981;

Polani, 1969b; Van Dyke, Weiss, Roberson, & Babu, 1983). These phenotypically normal individuals (carriers) are at high risk of producing chromosomally unbalanced gametes and embryos, resulting in much higher rates of recurrent miscarriage compared to those with numerical chromosome abnormalities (Keymolen, Staessen, Verpoest, Liebaers, & Bonduelle,

2012).

23 1.2.3. Origin of Chromosome Abnormalities in Human

There are three main stages at which numerical chromosome abnormalities may occur: gametogenesis including oogenesis and spermatogenesis, fertilization and embryogenesis

(Griffiths et al., 2000). Structural chromosome abnormalities are mostly inherited. However, the majority of numerical chromosome abnormalities are de novo, at which about 95% are of maternal origin, while around only 5% are paternally contributed.

Early analysis demonstrated that the predominance of these anomalies is maternally related

(Antonarakis, 1991; J. H. Ford et al., 1996; T. Hassold et al., 1984; T. J. Hassold, 1986; T. J.

Hassold & Jacobs, 1984; P. A. Jacobs, 1992; Martin et al., 1983; Martin et al., 1986). Further extensive cytogenetic analyses using different techniques on human oocytes and preimplantation embryos have confirmed that the most numerical chromosome abnormalities are attributed to errors that occur during oogenesis (Chatziparasidou et al., 2014; Delhanty & Handyside, 1995;

T. Hassold et al., 1996; T. Hassold et al., 2007; T. Hassold & Hunt, 2001; Templado et al.,

2013).

In the following section, the chromosomal complement and behaviour during the timeline of oogenesis in human will be described. Next, the different types of numerical chromosomal errors occurring during gametogenesis (oogenesis and spermatogenesis), fertilization, and embryogenesis will be discussed.

1.2.3.1. Chromosomal Complement During Human Oogenesis

Oogenesis is defined as the biological process of producing fully matured, competent female gametes (Baker, 1968; Gondos, Westergaard, & Byskov, 1986; Gougeon & Testart,

1986). In humans, oogenesis is enormously complex as compared to spermatogenesis (Figure

1.2).

24 Oogenesis in Humans

Oogenesis starts during the early stages of embryogenesis – at around 6-8 weeks gestation

(Gondos et al., 1986) as illustrated in Figure 1.3. Primordial germ cells (PGCs), which originate from the primitive streak and migrate to the gonadal ridge, start to proliferate via rapid and successive pre-meiotic mitotic divisions. These newly divided germ cells become sheathed with coelomic epithelial cells, which constitute the outermost layer of the female gonads after sex determination. At about 11-12 weeks of gestation, sexual differentiation is determined and the gonadal cords arise; the regulatory factors required for female sexual differentiation and the initiation of meiotic division remain poorly understood. During this period and up to around 22 weeks of gestation, PGCs in the foetal ovaries continue to proliferate mitotically, producing clones of oogonia. These oogonia remain connected with each other through cellular bridges due to incomplete cytokinesis. Once oogonial cells enter the prophase stage of the first meiotic division, they are called primary oocytes. Their number peaks to 7 million per ovary reach at approximately the 20th week of gestation, followed by a dramatic decline up to the 28th week of gestation,that is attributed to atresia. . These events give rise to an approximate total of 2 million primary oocytes per ovary at the time of birth. In fact, atresia starts concomitantly with oocyte proliferation at the 14th week of gestation. However, the proliferation rate during 14-20 weeks of gestation is significantly higher compared to rate of atresia, which steadily declines after the 28th week of gestation, giving rise to 400 oocytes in a woman's reproductive lifetime. The first pool of primary oocytes enters meiotic prophase at around the 11-12th week of gestation. Then, a quantitative increase in the number of oogonia enters the prophase stage up to the 25th week of gestation. The chromatin in these primary oocytes undergoes dramatic changes as it progresses through the sub-stages of prophase: Leptotene, Zygotene, and Pachytene (Figure 1.4). Within the

25 Diplotene stage, primary oocytes arrest for the first time at dictyotene. The arrested primary immature oocytes become surrounded by somatic cells called pre-granulosa cells, forming primordial follicles. In total, the time spent in the prophase stage in humans is approximately 4 weeks. Therefore, the first pool of primary oocytes that entered meiosis at the 11-12th week of gestation is expected to arrest until about the 15-16th week of gestation.

The arrested primary oocytes within the primordial follicles are maintained in their current state, and no signals to resume meiosis are given until the age of puberty (Gilbert, 2000). During each menstrual cycle, a cohort of primordial follicles is recruited for folliculogenesis and the primary oocytes are selected for growth and maturation. A highly specialized, complex interaction between the oocyte and the somatic cells of follicle is required to maintain good quality and developmentally competent oocytes ((R. Li & Albertini, 2013; Sobinoff, Sutherland,

& McLaughlin, 2013). These primary oocytes increase approximately 10-fold in size.

Concomitantly, an accumulation of cytoplasmic material is highly regulated for further successful fertilization and early embryonic development, including modifications in the subcellular energy-producing organelles, precursors for DNA, mRNAs, synthesized structural proteins and enzymes, and morphogenetic regulatory factors that control early embryogenesis

(Coticchio, Dal-Canto, Guglielmo, Mignini-Renzini, & Fadini, 2012) . On the other hand, primordial follicle development consists of several stromal cell divisions and differentiation following which these primordial follicles become primary, and subsequently, secondary follicles. Under the influence of Follicle Stimulating Hormone, secondary follicles develop to antral, and eventually to pre-ovulatory follicles. These immature oocytes, the so-called secondary oocytes, with fully developed cytoplasm within pre-ovulatory follicles are estimated to take about 85 days in humans to develop (Gougeon & Testart, 1986). In response to a Luteinizing

26 Hormone (LH) surge, only one secondary immature oocyte of the cohort resumes the first meiotic division at which the first reduction of chromosomal material is completed and the first polar body (1PB) is extruded. The first meiotic division then proceeds to the second meiotic division immediately, at which time, the oocyte arrests for the second time at the metaphase stage of the second meiotic division (MII). This mature oocyte is ovulated for fertilization or degeneration. If a successful fertilization occurs, the second meiotic division is triggered to resume when the second reduction of chromosomal material is completed and second polar body

(2PB) is extruded. The zygote with two pro-nuclei (2PN) undergoes DNA replication for the first post-zygotic mitotic division.

Spermatogenesis in Humans

In spermatogenesis, a brief period of mitotic proliferation takes place prior to sexual differentiation, followed by an extended period of mitotic arrest until the time of birth (Grinsted

& Byskov, 1981; Luciani, Devictor, & Stahl, 1977). After birth, spermatogonia resume mitotic proliferation, producing primary spermatocytes that undergo a further period of growth. Fully- grown primary spermatocytes are ready to initiate meiosis at the age of puberty. During meiosis, each primary spermatocyte undergoes the first meiotic division to produce two secondary spermatocytes. These secondary spermatocytes remain connected by cytoplasmic bridges. These connections are lost during post-meiotic divisions. Each of these secondary spermatocytes undergoes the second meiotic division escaping meiotic arrest to produce two spermatids, giving rise a total of four spermatids. Each of these spermatids develops into four fully matured spermatozoa (de Kretser, Loveland, Meinhardt, Simorangkir, & Wreford, 1998).

27

Figure 1.2: Oogenesis and spermatogenesis in humans. During foetal development, both female (top panel) and male (bottom panel) germ cells undergo successive proliferative mitotic divisions. In females, germ cells enter meiosis directly after the proliferative stage, followed by meiotic arrest prior to birth. A substantial number of the arrested oogonia undergo apoptosis, while the surviving primary oocytes enter an extended period of meiotic arrest until puberty, at which timeprimordial follicles are formed. At the onset of ovulation, a pool of primordial follicles with primary oocytes is stimulated to undergo growth and maturation. Only one follicle becomes fully developed for further ovulation; meanwhile, the oocyte is fully-grown, mature, and arrested following the first meiotic division. In males on the other hand, germ cells are arrested after a brief proliferative stage, followed by an extended period of mitotic arrest prior to birth. After birth, spermatogonia resume mitotic divisions for proliferation and enter meiosis by the time of puberty. At this time, meiosis is initiated and fully developed sperm are produced though spermiogenesis.

Adopted from Hassold and Hunt (2001).

28

Figure 1.3: A timeline of early oogenesis in humans. The first pool of germ cells undergoes proliferation at the 6-8th week of gestation; the first pools of oogonia are formed at the 9-11th week of gestation (B). Subsequently, primary oocytes enter the prophase stage at the 12-14th week stage (C), and same pools of primary oocytes become arrested at the dictyate stage between the 16-18th to the 29th weeks of gestation (D). At around the 14th week a quantitatively increased decline in the number of germ cells commences as well as atresia in all of the follicle stages (E). The number of germ cells undergoes a substantial variation in terms of total number due to the overlapped processes of proliferation and atresia. Germ cell numbers demonstrate substantial variability owing to different rates of proliferation and loss. The graph above depicts the number of germ cells across the lifespan of a female, ranging from the time of conception to 50 years of age. Between the 6th and 20th weeks of gestation, while germ cells undergo atresia, the rate of atresia is significantly lower than the rate of proliferation. In contrast, from the 20th week of gestation up to the time of birth, the rate of atresia is considerably higher, and the number of proliferative germ cells reaches their peak of about 7 million at the 20th week of gestation, following which the number declines dramatically to approximately 2 million at the time of birth.

Adopted from http://www.embryology.ch/anglais/cgametogen/oogenese03.html

29 Chromosome Configuration and Number during Oogenesis

Chromosomes behave differently during oogenesis (Figure 1.4), as their number is reduced during the reduction divisions of meiosis (Figure 1.5) (Table 1.3) (Baker, 1968; Gougeon &

Testart, 1986; Guichaoua et al., 1986).

Chromosomal configuration during the prophase I stage

Primary oocytes enter prophase as homologous chromosomes undergo pairing, synapsis, and crossing over for recombination. In the Leptotene sub-stage, also referred to as a the

Bouquet stage, homologous chromosomes have a thread-like appearance In the Zygotene stage, the homologous chromosomes’ pairing is initiated; a physical contact is generated between identical regions of the homologs. In the late Zygotene stage, total pairing is accomplished as synaptonemal complexes are formed between the paired homologous chromosomes; here, these fully paired chromosomes are referred to as bivalents. In the Pachytene stage, the bivalents begin to get shorter and thicker, and the interaction between them is strengthened by cohesins as they are fully synapsed. At this stage, the centromeres of each homologous chromosome are fully attached, and yet are not visually distinguishable. During synapsis, genetic material exchange between the two non-sister maternal and paternal chromatids per bivalent takes place (crossing over), resulting in recombinant chromosomes; these sites of recombination are called chiasmata.

In the Diplotene stage, the crossed over homologous chromosomes are separated and the majority of synaptonemal complexes and cohesins are lost. However they remain attached at the sites of the chiasmata and the cohesins around the centromeres and chromosomal arms. The centromeres of each homologous chromosome remain attached. Primary oocytes with these bivalents move on to the dictyate stage and are arrested. These bivalents are confined to the nuclear membrane, and the nucleus is enlarged giving rise to the distinct morphological

30 characteristics of the arrested primary oocytes. At this stage, they are referred to as immature oocytes at the germinal vesicle (GV) stage. These GV oocytes remain dormant at this stage until puberty.

Chromosomal complement during pre-meiotic, meiotic, and post-zygotic divisions

As shown in (Figure 1.5) (Table 1.3), diploid germ cells carry two copies of the chromosomes (2C) following each mitotic division during the proliferative stage in the foetal female gonads. Upon initiation of meiosis, the diploid oogonia carry two copies of the replicated chromosomes, each consisting of two sister chromatids (4C). At the time of arrest, the primary oocytes carry two copies of the recombined bivalents (4C) up to the time of puberty. At this point, the primary oocytes resume meiosis under the stimulation of Leutinizing Hormone (LH), and produce a diploid secondary oocyte (2C) and its corresponding diploid first polar body

(1PB). At the time of fertilization, the haploid secondary oocyte and its corresponding second polar body (2PB) carry one copy of maternal chromatids (1C). The resulting diploid zygote carries one copy each of the maternal chromatids (1C) and paternal chromatids (1C) and undergoes early embryogenesis, giving rise to diploid cells after each mitotic division.

On the other hand, in the male foetal gonads, the diploid germ cells carry two copies of the chromosomes (2C) after each pre-meiotic mitotic division during the proliferative stage. Meiosis is initiated in the foetal testes at puberty, and the diploid spermatogonia carry two copies of the replicated chromosomes (4C). After prophase I, the diploid primary spermatocytes carry two copies of the recombined bivalents (4C). The diploid secondary spermatocytes carry two copies of the chromosomes (2C) after the first meiotic division, and the haploid spermatids carry one copy of paternal chromatid (1C) after the consecutive second meiotic division. Fully developed haploid sperm (1C) are produced through spermatogenesis.

31

Figure 1.4: The chromosomal behaviour during the prophase stag in oogenesis. Chromosomal organization varies greatly during the Prophase versus proliferative stages of mitosis. Following DNA synthesis, chromosomes enter Prophase I, which is further subdivided into five stages: Leptotene, Zygotene, Pachytene, Diplotene, and Dictyate, each containing varying chromosomal configuration. The top panel illustrates chromosomal organization during the four sub-stages of Prophase I while the bottom panel depicts chromosomal organization during oogenesis.

Adopted and modified from Homer (2013).

32

Figure 1.5: The chromosomal complements during pre-meiotic mitotic, meiotic and post- zygotic mitotic divisions in human. During the proliferative stage in the foetal female gonads, (i) diploid germ cells carry two copies of the chromosomes (2C) following each mitotic division. Upon the initiation of meiosis in the foetal ovaries, (ii) the diploid oogonia carry two copies of the replicated chromosomes, each consisting of two sister chromatids (4C). Following meiotic arrest at Prophase I, the diploid primary oocytes carry two copies of the recombined bivalents (4C). Later, at puberty, the primary oocytes resume meiosis under the stimulation of Luteinizing Hormone (LH), and produce a diploid secondary oocyte (2C) and its corresponding diploid first polar body (1PB).. At the time of fertilization, the haploid secondary oocyte and its corresponding second polar body (2PB) carry one copy of maternal chromatids (1C) (iii) The resulting diploid zygote carries one copy each of the maternal chromatids (1C) and paternal chromatids (1C) and undergoes early embryogenesis, giving rise to diploid cells after each mitotic division. During the proliferative stage in foetal male gonads (iv), diploid germ cells carry two copies of the chromosomes (2C) after each mitotic (pre-meiotic) division. In contrast to females, meiosis is initiated in the foetal testes at puberty (v), and the diploid spermatogonia carry two copies of the replicated chromosomes (4C). After prophase I, the diploid primary spermatocytes carry two copies of the recombined bivalents (4C). The diploid secondary spermatocytes carry two copies of the chromosomes (2C) after the first meiotic division, and the haploid spermatids carry one copy of paternal chromatid (1C) after the consecutive second meiotic division. Fully developed haploid sperm (1C) are produced though spermatogenesis.

Adopted from Jones and Lane (2013).

33 Table 1.3: The Chromosomal Complements in Germ Cells and Human Oocytes at Different stage. Cell Type Chromosomal Complement Meiotic Stage Biological stage Premordial Germ Cells Germ Cells Diploid (2N/2C, N=46) Pre-meiotic Oocytogenesis (PGCs) Oogonia Germ Cells Diploid (2N/2C, N=46) Pre-meiotic Oocytogenesis Primary Oocytes Immature Oocytes Diploid (2N/4C, N=46) Prophase I Ootidogenesis and Folliculogenesis Secondary Oocytes Mature Oocytes Diploid (2N/2C, N=46) Metaphase II Ootidogenesis Zygote Fertilized, two-pronuclear Haploid (1N/1C, N=23) Post-meiotic Fertilization Cell (2PN)

34 Here, the different types of chromosomal errors will be discussed as chronologically occurred during oogenesis to early embryogenesis.

1.2.3.2. Pre-Meiotic Mitotic Errors

Chromosome abnormalities may arise during oogenesis prior to meiotic divisions as pre- meiotic mitotic errors.

Hypotheses of The Origin of Pre-meiotic Mitotic Errors

Two hypotheses have been proposed to explain the origin of pre-meiotic mitotic errors, as reviewed by (Biesecker & Spinner, 2013; Delhanty, 2011; Papavassiliou et al., 2015; Taylor et al., 2014). The first hypothesis proposes that these pre-meiotic mitotic errors may arise as a result of mitotic errors occurring during germ cell proliferation in which chromosomally abnormal cells may ultimately become gametes. The second hypothesis suggests that pre-meiotic mitotic errors may arise during pre-implantation embryonic development in which chromosomally abnormal cells may eventually become precursors to germ cells. These pre-meiotic mitotic errors have been defined as germline/gonadal mosaicism. Both hypotheses are illustrated in Figure 1.1.

Underlying Mechanisms Leading to Pre-meiotic Mitotic Errors

Non-disjunction (ND) has been proposed as the main mechanism leading to pre-meiotic mitotic errors during the successive proliferative stage. Therefore, different chromosomes might be involved in the chromosomally abnormal germ cells due to pre-meiotic mitotic errors (Cupisti et al., 2003; Daina et al., 2014; Mahmood et al., 2000; Morris et al., 2012; Obradors et al., 2010;

Pujol et al., 2003; Rowsey et al., 2013). However, it has been proposed that same chromosome can be involved in the resulting abnormal germ cells which are generated from mitotic errors occurred during preimplantation embryonic development of the same individual (Cozzi et al.,

1999; Mahmood et al., 2000; Rowsey et al., 2013). Different leading mechanisms to these

35 mitotic errors in preimplantation embryos have been well documented, including ND and anaphase lag (AL). These mechanisms will be defined and discussed in the subsection post- zygotic errors, as reviewed by (Mantikou et al., 2012; Taylor et al., 2014). Accordingly,

Delhanty (2011) suggested that the terminology of germline mosaicism is preferred to define those errors arise prior to meiosis during the proliferative stage, while gonadal mosaicism is reserved to errors arise during the preimplantation embryonic development.

The Overall Rate of Pre-meiotic Mitotic Errors

To date, the contribution of pre-meiotic mitotic errors to the overall rate of chromosome abnormalities in the general population remain unknown. Limited studies have been conducted to investigate the prevalence of pre-meiotic mitotic errors in human foetal oocytes obtained from women with normal karyotypes; all these studies are listed in Table 1.2-A. Conflicting results were reported by these studies. Only two studies were designed to analyze the prevalence of pre- meiotic mitotic errors in human adult oocytes retrieved from women who underwent ART treatment (Table 1.1-B-C) (Daina et al., 2014; Obradors et al., 2010). Also, inconsistent results were described by both studies.

Observations from limited studies show a great discrepancy in the overall rate of pre- meiotic mitotic errors in human mature oocytes with the corresponding first polar bodies derived from women who underwent ART treatment cycles. This is largely because these studies were not intended to investigate this type of chromosomal error (Table 1.2-B). Most of these studies were designed to develop and standardize a larger panel of FISH probes or CGH techniques for chromosomal abnormality screening in mature oocytes and polar bodies. A high rate of experimental artifacts, as well as the small sample size, mixed population of oocytes (unfertilized oocytes and in vitro matured oocytes which were not subjected to sperms), and the diverse

36 population of female subjects (donors or patients with fertility problems), are all contributing factors to the incidence of pre-meiotic mitotic errors.

Although there was considerable heterogeneity in the investigated overall rate of these errors, all these studies conducted results concluded that the overall incidence of these pre- meiotic mitotic errors is relatively low, and germ cells carried an extra chromosome due to pre- meiotic mitotic errors can reach the stage of immature and mature oocytes.

The Extra Chromosome Behaviour During Prophase I Stage To Anaphase I Stage

The alignment and paring of the extra chromosome during the prophase stage, and the subsequent possible segregation patterns during anaphase I stage have been investigated in human oocytes (Barlow & Hulten, 1998; Barlow, Tease, & Hulten, 2002; E. Y. Cheng, Chen,

Bonnet, & Gartler, 1998; E. Y. Cheng, Chen, Disteche, & Gartler, 1999; E. Y. Cheng, Chen, &

Gartler, 1995; E. Y. Cheng & Naluai-Cecchini, 2004; Y. E. Cheng & Gartler, 1994; Cozzi et al.,

1999; Cupisti et al., 2003; Hulten et al., 2008; Lenzi et al., 2005; Luciani, Devictor, Morazzani,

& Stahl, 1976; Mahmood et al., 2000; Prieto et al., 2004; Pujol et al., 2003; Robles et al., 2007;

Roig, Robles, Garcia, Martin, et al., 2005; Roig, Robles, Garcia, Martinez-Flores, et al., 2005;

Speed, 1984, 1985; Tease, Hartshorne, & Hulten, 2006; Tease, Hartshorne, & Hulten, 2002;

Wallace & Hulten, 1983). Most of these studies suggested that the presence of an extra copy of chromosome does not affect the cohesins uploading and organization (Prieto et al., 2004) and the synaptonemal complex (SC) formation for subsequent pairing and synapsis (Barlow et al., 2002;

Luciani et al., 1976; Roig, Robles, Garcia, Martin, et al., 2005; Roig, Robles, Garcia, Martinez-

Flores, et al., 2005). Same studies as well as others showed a higher incidence of trivalent formation of the extra chromosome than bivalent with univalent for chromosomes 13, 16, 18, and 21(Hulten et al., 2008; Lenzi et al., 2005; Robles et al., 2007; Speed, 1984, 1985; Tease et

37 al., 2006; Tease et al., 2002; Wallace & Hulten, 1983). This suggests that the trivalent formed at the Pachytene stage can be maintained until the Diplotene stage, although the exact mechanism by which these homologues interact and recombine is not clear.

The subsequent possible segregation patterns of the extra chromosome during anaphase I stage have been proposed by (Hulten et al., 2008) (Figure 1.6). Both patterns have been observed in human mature oocytes with the corresponding 1PBs (Cozzi et al., 1999; Cupisti et al., 2003;

Fragouli, Wells, Whalley, et al., 2006; Gutierrez-Mateo, Benet, et al., 2004; Mahmood et al.,

2000; Obradors et al., 2010; Pujol et al., 2003). The relevant incidence of either patterns of the extra chromosome in human mature oocytes remains unknown. In the case of the extra chromosome maintained in a form of trivalent in the immature oocyte, the trivalent aligned at the metaphase plate at metaphase I stage as shown in the Figure 1.6. Subsequently, a ND occurs at anaphase I stage where either mature oocyte or the corresponding 1PB will receive the extra copy of the chromosome. Therefore, hypothetically, there is a probability that about 25% of the resulting mature oocytes will be aneuploid due to pre-meiotic mitotic errors. In contrast, in the case of the extra chromosome maintained in a form of bivalent and univalent in the immature oocyte, the bivalent and univalent are aligned at the metaphase plate at metaphase I stage separately as displayed in the Figure 1.6. Subsequently, a ND occurs at anaphase I stage where the bivalent segregates similarly to the rest into its homologues chromosomes, while the univalent segregate into its constituents chromatids by which each of mature oocyte and its corresponding 1PB will receive one copy of the chromatid. Therefore, hypothetically, the resulting mature oocytes will be all aneuploid due to pre-meiotic mitotic errors.

38

Figure 1.6: Proposed models for the alignment and segregation of a trisomic immature oocyte. (A) At prophase, the extra chromosome of a trisomic oocyte is aligned separately as a univalent, while the other two homologous of the same chromosome are aligned and paired together as a bivalent. Later, at metaphase I, the sister kinetochore of each homologous chromosome of the bivalent acts as a single kinetochore for amphitelic microtubule attachments and further segregation. The sister kinetochores of the extra univalent acts as two kinetochores for amphitelic microtubule attachment and further segregation. The resulting mature oocyte and its corresponding polar body thus consist of a homologous chromosome and an extra chromatid. (B) At prophase, the extra chromosome and the other two homologs of the same chromosomes are aligned and paired together as a trivalent. At metaphase I, the sister kinetochores per homologous chromosome of the trivalent act as one for amphitelic microtubules attachments and segregation. The resulting mature oocyte and its corresponding polar body consists of one and two homologous chromosomes, respectively, or vice versa.

Adopted from Hulten et. al. (2008).

39 Spindle assembly checkpoint (SAC) has been proposed to play a role in the possibility of immature oocytes carried an extra chromosome due to pre-meiotic mitotic errors to undergo the first meiotic division for maturation (Delhanty, 2011). Normally, SAC is activated whenever a single chromosome is unattached to the spindles and a delaying in anaphase I occurs, as has been recently reviewed by (Jones & Lane, 2013). This suggests that a bi-orientated attachment of the extra univalent chromosome as well as the bivalent at metaphase I stage of the chromosomally abnormal immature oocyte may avoid activating the SAC pathway (Kouznetsova, Lister,

Nordenskjold, Herbert, & Hoog, 2007). Thereby, the univalent segregates prematurely at anaphase I stage, and each chromatid will be involved in the nuclei of mature oocytes and its

1PB. Similarly, it has been proposed that a bi-orientated attachment of the trivalent may develop, resulting in deactivation of the SAC mechanism (Garcia-Cruz, Casanovas, et al., 2010; Hulten et al., 2010).

Pre-meiotic Mitotic Errors in Relation to Maternal Age

A model called the Oocyte Mosaicism Selection Model (OMSM) has been proposed to explain the maternal age effect on the increased rate of chromosome abnormalities in human

(Hulten et al., 2010). This model suggests that pre-meiotic mitotic errors are maternal-age dependent; a selection against the chromosomally abnormal immature oocytes for ovulation might occur during the female reproductive lifetime, resulting in an increase of those oocytes within the antral follicle count among women of advanced age compared to younger women.

Only one study has been conducted to investigate the incidence of pre-meiotic mitotic errors in human immature oocytes in relation to maternal age (Daina et al., 2014). The results showed no association between pre-meiotic mitotic errors in immature oocytes and the advanced maternal age. Further studies are required to examine the model in human immature and mature oocytes.

40 1.2.3.3. Meiotic Errors

Chromosome abnormalities may also arise as meiotic errors during oogenesis; these errors can occur at first meiotic division (first meiotic errors MI) and or occur at second meiotic division (second meiotic errors MII).

Underlying Mechanisms Leading to Meiotic Errors

There are three main mechanisms leading to meiotic errors inhuman oocytes: ND, anaphase lagging (AL), and premature division (PD) (Figure 1.7) (Griffiths et al., 2000). ND and

AL has been known as the classical and main leading mechanisms to meiotic errors in human oocytes (Kuliev & Verlinsky, 2004). However, PD is well known now as the most predominant mechanism leading to meiotic errors (Capalbo et al., 2013; Gabriel et al., 2011; Handyside et al.,

2012; Kuliev et al., 2011).

The first mechanism, ND, is described as the failure of homologous chromosome to be segregated to the opposite poles in a typical amphitilic manner during the anaphase I stage.

Instead, in ND at the first meiotic division, the homologous chromosomes of a bivalent are pulled to either pole in a syntelic fashion, resulting in nullisomic (2N-1) or disomic (2N+1) oocyte for the same chromosome involved. In ND at the second meiotic division, the sister chromatids of a homologous chromosome are segregated to either pole during anaphase II stage, resulting monosomic (N-1) or trisomic (N+1) zygote of the same chromosome involved, fertilized by a chromosomally normal sperm (N). This syntelic segregation occurs because of the attachments of both pairs of sister kinetochores to the same spindle pole. This results in lacking of equal tension generation along the bivalent or homologous in first and second meiotic divisions, respectively, and thus moving to either pole. The second mechanism, AL, is defined as the failure of any of the homologous chromosomes to be pulled after segregation to the

41 opposite poles during anaphase I stage. In contrast to ND, in AL, one of the kinetochore pairs of bivalent attaches to spindles of opposite poles, while the other pair remains attached to either pole. The unequal pulling forces on the former pair of kinetochores, resulting in homologue lagging, instead of being pulled towards one pole. This type of segregation called merotelic. As a result to this faulty attachment, a monosomic (2N-1) or euploid (2N) oocyte of the same chromosome involved. Whereas, AL at the second meiotic division is known as the failure of any of the sister chromatids to be pulled to the opposite poles during anaphase I stage due a similar faulty mechanism described previously in the first meiotic division. Subsequently, AL may result in monosomic (N-1) or euploid (N) zygote of the same chromosome involved, fertilized by a chromosomally normal sperm (N) (Delhanty, 2005). The last mechanism found to be leading to meiotic errors in human oocytes is PD, or premature centromere division (PCD), or premature separation of sister chromatids (PSSC). In PD, the sister chromatids are separated from each other during anaphase I (AI) due to a premature division of the sister chromatids, resulting in metrotelic segregation of these chromatids. In contrast to AL, in PD, equal pulling forces on the pair of sister kinetochores, resulting in complete chromatid pulling to the opposite pole. Therefore, this type of mechanism may result in a mature oocyte with an additional (2N+½) or missing (2N-½) chromatid. Subsequently, these numerically unbalanced oocytes may result in trisomic (N+1) or monosomic (N-1) zygotes of the same chromosome involved, fertilized by a chromosomally normal sperm (N). PD may also result in an euploid oocytes (2N), numerically balanced oocytes, where both separated chromatids are moved to the opposite poles. However, these oocytes are at risk of 50% to produce chromosomally abnormal zygote for the same chromosome involved.

42 First meiotic errors resulted by PD has been found to be strongly associated with advanced maternal age compared to errors resulted by ND (Gabriel et al., 2011; Kuliev et al., 2011). The underlying molecular basis for PD leading mechanism to first meiotic errors in association to maternal age has been studied recently (Chiang, Schultz, & Lampson, 2012; Jessberger, 2012); this will be discussed in the last section of the present review. In contrast to PD and ND, AL leading to meiotic errors has not been found to be in association with maternal age (Magli et al.,

2012).

PD has been proposed as a possible mechanism that may contribute to the MI errors after analyzing unfertilized oocytes (Angell, 1991). Since then, many cytological and molecular studies using different techniques have supported Angell’s observation (Chiang, Duncan,

Schindler, Schultz, & Lampson, 2010; Gabriel et al., 2011; Garcia-Cruz, Brieno, et al., 2010;

Handyside et al., 2012; Kuliev et al., 2011; Magli et al., 2012; Velilla, Fernandez, Sunol, &

Lopez-Teijon, 2013). All studies have concluded that PD is, in fact, the most predominant mechanism leading to meiotic I (MI) errors during oogenesis, followed by AL, and ND is the least prevalent mechanism. Only ND occurring at first meiotic division lead to abnormal oocytes and resulting embryos in almost all cases; however, ND occurring in second meiotic division, PD occurred in first meiotic division, and AL in either meiotic division will lead to abnormal oocytes and resulting embryos in about 50% of cases. All related clinical and research studies on meiotic errors in human oocytes are summarized in (Table 1.1).

43

Figure 1.7: Proposed mechanisms leading to first and/or second meiotic errors in human mature oocytes. (A) Schematic of normal meiotic divisions leading to a euploid zygote and its corresponding polar bodies. (B) Abnormal first and/or second meiotic divisions leading to aneuploid mature oocytes, zygotes, and/or their corresponding polar bodies; (i-ii) Classic non-disjunction leading to meiotic errors at first and/or second meiotic divisions; (iii-iv) Premature division leading to meiotic errors at first and/or second meiotic divisions; and (v-iv) Anaphase lag leading to meiotic errors at first and/or second meiotic divisions.

Adopted and modified from Jones and Lane (2013).

44 Meiotic Errors in Relation to Maternal Age

Meiotic errors have been well documented to be strongly linked to maternal age; overall rate of meiotic errors dramatically increase in women of their late 30s (T. J. Hassold & Jacobs,

1984). However, the full picture of underlying molecular basis of the effect of advanced maternal age remains unclear (Jones & Lane, 2013). Many hypotheses have been proposed in order to elucidate the advanced maternal age effect on the increased rate of chromosome abnormalities.

Most of the postulated hypotheses are based on any faulty event arising during the developmental stage of oogenesis; these hypotheses are reviewed in (Rowsey et al., 2013).

Among of these hypotheses, two models which have been studied extensively: the complete production line model; and the prolonged meiotic arrest model.

The production line model is the oldest postulated hypothesis after observing a significant increase in the frequency of univalents in mature oocytes derived from old mice compared to those derived from young mice (Henderson & Edwards, 1968). Henderson and his colleague suggested that the increased rate of univalents is maternally age related. These univalents are attributed to bivalents without any chiasmata (achismatic), or bivalents with a single chiasma that either distal or proximal to the centromere in the germ cells. They postulated that since recombination takes place during early oogenesis in the foetal ovaries, the last pool of oogonia to enter meiosis are the last to be ovulated as secondary oocytes, which undergo inadequate recombination. Therefore, last oocytes to be ovulated are the most susceptible to nondisjunction.

Many observations have been reported in order to examine the complete production line model; however, only conflicting results have been obtained (Coop, Wen, Ober, Pritchard, &

Przeworski, 2008; Hussin, Roy-Gagnon, Gendron, Andelfinger, & Awadalla, 2011; Kong et al.,

2004; Polani & Crolla, 1991; Rowsey, Gruhn, Broman, Hunt, & Hassold, 2014). Therefore, the

45 exact underlying molecular mechanisms of the maternal age effect for this model remain poorly understood.

The meiotic arrest model is the most favourable hypothesis after discovering the significant depletion of cohesions in mature oocytes derived from old mice compared to those derived from young mice; similar findings has been reported in human oocytes in relation to maternal age

(Chiang et al., 2010; Hodges, Revenkova, Jessberger, Hassold, & Hunt, 2005; Lister et al., 2010;

L. Liu & Keefe, 2008; Tachibana-Konwalski et al., 2010; Tsutsumi et al., 2014). It has been found that the cohesin complexes in meiosis has dual functions in binding the homologue around the distal arm regions, and the sister chromatids around the centromere regions for proper segregation at first and second meiotic divisions, respectively. Although these complexes are loaded during the DNA synthesis at early foetal development, cohesions around the centromere regions are found to be more degraded in oocytes derived from older women compared to its counterparts derived from younger women. Moreover, the level of Rec8 and Smc1 have been found to be decreased significantly in women at their 40s compared to those at their 20s

(Tsutsumi et al., 2014). Such depletion contributes to the premature separation of homologs and/or sister chromatids, resulting in first meiotic errors.

1.2.3.4. Fertilization Errors

Another stage where chromosome abnormalities may arise is around fertilization, resulting in hyperploid zygotes (tripronuclear zygotes (3PN) or hypoploid zygotes (mononuclear zygote

(1PN)). The overall rate of these hyperploid and hypoploid zygotes has been observed to be ranged from 2% to over 30% and 10% up to 50%, respectively (Egozcue, Blanco, Vidal, &

Egozcue, 2002; Feng & Hershlag, 2003; Munne, Marquez, Reing, Garrisi, & Alikani, 1998).

46 Human hypoploid zygotes containing Y chromosome has been observed after insemination and

ICSI. Embryos generated from these zygotes after insemination showed mostly diploid status and achieved successful pregnancy. This has been explained by the inability to observe both pronuclear or to catch both pronuclear before the fusion. However, only poor quality, arrested embryos generated from these zygotes after ICSI have been observed. It has been suggested that zygotes of these embryos to be parthenogenetically activated not fertilized (Munne et al., 1998;

G. D. Palermo, Munne, Colombero, Cohen, & Rosenwaks, 1995). This parthenogenetic activation has been seen after ICSI as well, which may occur either spontaneously or secondary to the injection procedure. It has been suggested that parthenogenetic activation post ICSI may linked to the calcium content of the injection medium and to the postovulatory age of the oocytes

(Feng & Hershlag, 2003).

On the other hand, Hyperploidy of human zygotes have been observed to be generated at three different events; first, fertilization by diploid sperm, which is called dispermy; second, fertilization by diploid oocyte, which is called digyny; and third, inhibition of second polar body extrusion and formation of two female pronuclei with a single sperm pronucleus, which is called monospermic digyny. In the first two scenarios, digyny and dispermy are distinguished morphologically by the presence of number of extruded polar bodies. Human tripronuclear zygotes have a remarkable clinical significance because it is known that these zygotes contribute to spontaneous abortions (Feng & Hershlag, 2003; G. D. Palermo et al., 1995). It has been observed that about half of the triploid zygotes reached the 4–6-cell stage of development with multinuclear cells. Less than 10% of these examined zygotes even reached early morulas or blastocyst stage (Balakier, MacLusky, & Casper, 1993). It has been reported also that embryos

47 developed from dispermic zygotes are mostly mosaics (diploid/triploid), around 60%, compared to embryos developed from digynic zygotes are triploids (Munne et al., 1998).

Errors Around the Time of Fertilization in Relation to Maternal Age

Errors at fertilization have not been found to be associated with advanced maternal age

(Feng & Hershlag, 2003).

1.2.3.5. Post Zygotic Errors

As chromosome abnormalities can occur as pre-meiotic mitotic, meiotic errors, and fertilization errors, numerical chromosome anomalies may also arise as post-zygotic mitotic errors during developmental stage of preimplantation embryos: form two-cell embryos to blastocyst stage.

Underlying Mechanisms Leading to Post-Zygotic Mitotic Errors

There are two main mechanisms leading to mitotic errors in human preimplantation embryos: anaphase lagging (AL) and mitotic non-disjunction (ND) (Griffiths et al., 2000). AL during mitosis is described as the failure of the sister chromatids to be pulled to the opposite poles during anaphase stage. These lagging chromatids are lost and, subsequently, are not incorporated in either nucleus of the divided cells. This resulting in a monosomic cell for that chromosome involved, while the other cell remains diploid for the same corresponding chromosome. The other classical mechanism is mitotic ND which is defined as the failure of the sister chromatids to be separated to the opposite poles during anaphase stage. These non- disjunction chromatids, subsequently, are incorporated in either nucleus of divided cells. This resulting in one monosomic cell for the chromosome involved, while the other cell is trisomic for the same chromosome involved.

Several studies demonstrated that AL is the major mechanism leading to chromosomal

48 mosaicism in human preimplantation embryos compared to the classical ND (Capalbo et al.,

2013; Chow et al., 2014; Coonen et al., 2004; Daphnis et al., 2005; Ioannou et al., 2012). Some of these studies reported a higher rate of monosomic cells of about 8 times (Daphnis et al., 2005),

7 times (Ioannou et al., 2012), and 5 times (Coonen et al., 2004) higher than trisomic cells in total diploid-aneuploid mosaic embryos using a 3-probe, 24-probe and 5-probe FISH panels, respectively. Observations from other studies using aCGH supported that AL is the predominant mechanism leading to chromosomal mosaicism in diploid-aneuploid mosaic embryos, while ND is the common mechanism leading to mosaicism in aneuploid-chaotic mosaic embryos (Capalbo et al., 2013; Chow et al., 2014). However, many cytogenetic studies highly recommended that monosomies should be interpreted with an extra caution when FISH is used as a technique for chromosomal complements analysis. A monosomic signal could be interpreted as a false positive due to hybridization failure or overlapping signals (Fragouli, Alfarawati, Daphnis, et al., 2011).

An incident of monosomy could be due to ND combined with other mechanisms resulting in losing one of the trisomic signals or just lost due to a hybridization failure (Mantikou et al.,

2012).

The other mechanisms leading to mitotic errors have been proposed in human preimplantation embryos are: premature cell division, endoreplication, cell fusion, errors in cytokinesis, chromosome demolition, and chromosome breakage. These mechanisms have been observed in very limited studies which were found to be in association with embryos obtained from abnormally fertilized or parthenogenatically activated zygotes, embryos were frozen- thawed, multinucleated, or arrested, and embryos with structural aberrations. Therefore, the frequency rate of these mechanisms remains significantly lower in these limited studies compared to AL and ND (Mantikou et al., 2012; Taylor et al., 2014).

49 In premature cell division, the chromosomes are segregated without a replication occurring during the mitotic division; this results in haploid cells. In contrast, endoreplication, cell fusion, and errors in cytokinesis result in polyploid cells. In endoreplication, the chromosomes are replicated without mitotic division occurring. In cell fusion, both cells in a preimplantation embryo are fused; whereas, two cells are failed to divide or asymmetrically divided during the faulty cytokinesis. Haploidy was observed as uniformly haploid nuclei in multinucleated blastomeres per embryo with a significantly low rate compared to diploid nuclei. This has been suggested to maybe due to a combination of premature karyokinesis but not cytokinesis from a diploid mononucleated blastomere (Mantikou et al., 2012; Yilmaz et al., 2014). Haploidy was also observed as haploid cells in diploid-haploid mosaic embryos with two times higher rate compared to diploid-aneuploid mosaic (Mateo et al., 2013); however, these embryo were originated from monopronucleated (1PN) zygotes after ICSI. On the other hand, polyploidy was observed more frequently at which endoreplication has been suggested as the leading mechanism. For example, polyploidy was detected in arrested human zygotes (B. Rosenbusch,

Glaeser, Brucker, & Schneider, 2002; B. Rosenbusch, Schneider, & Sterzik, 1997) arrested and developed 2-4 cell, mononucleated embryos (Bielanska et al., 2002a), mononucleated trophectoderm in blastocysts (Bielanska et al., 2002b), and trophoblasts in post-implantation human placenta during the first trimester (Sarto, Stubblefield, & Therman, 1982).Eerrors in cytokinesis, however, has been suggested as a possible leading mechanism to polyploidy that was detected in bi or multinucleated embryos (Hardy, Winston, & Handyside, 1993; Yilmaz et al., 2014). On the other hand, cell fusion has been proposed as the underlying mechanism to polyploidy that was observed in frozen-thawed embryos, and the least in frequently among the previously mentioned mechanisms (Balakier et al., 2000).

50 Chromosome demolition and chromosome breakage are another possible mechanisms that may contribute to mitotic errors in human preimplantation embryos. Chromosome demolition involves a chromosomal destruction and cellular fragmentation, resulting in a monosomic cell for the chromosome involved (Mantikou et al., 2012). Chromosome demolition, in fact, has been suggested as a possible mechanism for the chromosomal rescue of trisomic cells (self-correction)

(Liehr, 2010; Los et al., 1998). On the other hand, chromosome breakage involves a partial chromosome breakage resulting in a partial monosomic cell for the same contributing chromosome (Mantikou et al., 2012). Tis mechanism has been proposed as the underlying mechanism for chromosomal instability (Vanneste et al., 2009), as well as de novo structural rearrangements via interstitial or terminal DNA double-stranded breaks and fusions in the zygote or during the first three cleavages (Voet et al., 2011). The frequency rate for all these leading mechanisms to mitotic errors including AL and ND have yet to be confirmed by systemic review and meta-analysis (Mantikou et al., 2012).

Underlying Molecular Mechanisms of Post-Zygotic Mitotic Errors

The underlying molecular mechanisms causing mitotic errors in human preimplantation embryos could be the results of maternal, paternal, and or others relevant factors. However, all following proposed molecular basis have yet to be confirmed by more investigations (Mantikou et al., 2012). Because the embryonic genome is activated at the 4-8 cell stage, the first mitotic divisions and chromosomal segregation are controlled by the oocyte mRNA and proteins.

Therefore, a defective in any of this maternal pool lead to faulty mechanisms related to chromosomal segregation and cell division. Microtubules, cell cycle checkpoints, cohesions, and

DNA repair proteins are all maternally inherited in the first mitotic divisions. For example, cell cycle checkpoints including SAC mechanisms safeguard correct mitotic cell divisions and

51 normal cell cycle at every cell cycle stage before progression to the next stage: G1, G2, and M stage (Decordier, Cundari, & Kirsch-Volders, 2008). The occurrence of any error will generate signals to cell cycle checkpoint mechanism which recruit the required proteins to halt cell division until repairs are completed. If the repair is not possible, then the checkpoint directs the cell towards apoptosis or arrest. RB and WEE1 are the key proteins of G1 and G2 cell cycle checkpoints, respectively. And similarly to meiosis, BUB3, BUBR1 and Mad2 are the key proteins of SAC in mitosis. Los of function of the later proteins has found to accelerate the metaphase-anaphase stage during the first cleavage stage resulting in micronuclei, and chromosome abnormality (Wei et al., 2011).

On the other hand, centrosome that is the core of establishing and organizing the mitotic spindle is paternally inherited (Carrell, 2008). Abnormalities in the number or kinetics of centrioles may result in abnormal spindle formation and chromosome mal-segregation during embryo development. It has been observed a higher rate of mosaic embryos due to abnormal spindle organization (G. Palermo, Munne, & Cohen, 1994).

Non-parental factors such as temperature fluctuation (Hong, Lee, Forman, Upham, &

Scott, 2014), oxygen concentration (Guo et al., 2014), prolonged in vitro culture (Zhang et al.,

2010), and culture medium in in vitro-generated embryos (Thrasher & Kilburn, 2001), as well as hormonal stimulation regimes (Rubio et al., 2010) have been reported to may associate with a higher rate of chromosome abnormalities in preimplantation embryos.

Post-Zygotic Mitotic Errors in Relation to Maternal Age

Post-Zygotic Mitotic Errors have not been found to be associated with advanced maternal age after the systemic review and meta-analysis performed in human preimplantation embryos

(Mantikou et al., 2012; van Echten-Arends et al., 2011).

52 Uniparental Disomy (UPD)

Uniparental disomy (UPD) can arise as a result to errors occurring at pre-meiotic mitotic

(Delhanty, 2011), meiotic (Jones & Lane, 2013), prost-zygotic mitotic divisions (Taylor et al.,

2014), or at fertilization (Golubovsky, 2003). UPD is well acknowledged phenomenon in which two homologous chromosomes (heterodisomy) or identical (isodisomy) chromosomes are inherited from either parent instead of one chromosome from each parent. UPD is linked to some serious recessive disorders or imprinting disorders. The prevalence of UPD in live births is estimated to be around 0.03% (Robinson, 2000). Similar prevalence has been found in human blastocysts, of about 0.06% (Gueye et al., 2014), while the overall rate of UPD in human cleavage stage embryos is estimated to be about 10-70% (Capalbo et al., 2013; Wells,

Alfarawati, & Fragouli, 2008), and around 3- 18% in human gametes (Robinson, 2000).

All previously mentioned mechanisms leading to meiotic and mitotic errors may contribute to UPD (Taylor et al., 2014), as well as other mechanisms which are reviewed in (Robinson,

2000). Chromosome 15 is the most involved chromosome in UPD (UPD15) that is responsible for Angelman syndrome (two paternally inheretied copies) or Prader-Willi syndrome (two maternally inherited copies) (Robinson, 2000).

53

CHAPTER TWO:

MATERIAL AND METHODS

54 2.1. FISH Analysis Standardization and Validation (Pre-examination Stage)

FISH-based analysis in the present project was validated (pre-examination process) prior to the experimental stage (examination and post-examination process) as recommended by the

ESHRE PGD consortium of the best practice guidelines for FISH-based PGD (Harton et al.,

2011).

2.1.1. FISH Probe Mixtures Preparation

Commercially available FISH probes for chromosomes 13, 15, 16, 18, 21, 22 and X

(Vysis, Downers Grove, IL, USA) were selected and prepared as previously described (Zhang et al., 2010). Two mixtures of FISH probes were prepared for two successive hybridization rounds.

Both FISH probe mixtures were then validated using patients’ lymphocytes and donated spare embryos, and only validated probe mixtures were applied on the experimental samples.

The first FISH probe mixture, a multicolour PB probe (MultiVysion PB probe, Vysis,

Downers Grove, IL, USA) for chromosomes 13 (Vysis LSI 13, Locus 13q14 in spectrum red), 16

(Vysis CEP 16, Locus 16q11.2 Satellite II DNA in spectrum Aqua), 18 (Vysis CEP 18, Locus

18p11.1-q11.1 Alpha Satellite DNA in spectrum Blue), 21 (Vysis LSI 21, Locus 21q22.13-q22.2 in spectrum green), and 22 (Vysis LSI 22, Locus 22q11.2 in spectrum orange) was diluted with hybridization buffer (CEP hybridization buffer, Vysis, Downers Grove, IL, USA) in a ratio of

1:1 as recommended by the manufacturer’s manual.

The second FISH probe mixture, a 2-color hybridization mixture specific to chromosomes

15 and X was prepared by combining the individual probes for chromosome 15 (Vysis CEP 15

D15Z1, Locus 15p11.2 Satellite III DNA in spectrum green) and chromosome X (Vysis CEP X

DXZ1, Locus Xp11.1-q11.1 alpha Satellite DNA) in a ratio of 1:1 as recommended by the manufacturer’s manual in hybridization buffer (CEP hybridization buffer, Vysis, Downers

55 Grove, IL, USA). The first and second FISH probe mixtures were used in the first and second rounds, respectively for the FISH analysis.

2.1.2. FISH Probes Mixtures Validation

FISH probe mixtures validation was performed using interphase nuclei from peripheral blood lymphocytes and interphase nuclei from donated spare human embryos in order to assess the hybridization efficiency before applying them on the experimental samples (Harton et al.,

2011).

The peripheral blood samples were withdrawn from patients with normal karyotype; the isolated peripheral blood lymphocytes were used for FISH probes validation as a normal

(negative) control. The donated spare embryos were obtained from patients underwent PGD cycles for translocation or aneuploidy screening; embryos diagnosed as abnormal were used for

FISH probes validation as an abnormal (positive) control.

A minimum of 100 interphase nuclei from peripheral blood lymphocytes and 50 interphase nuclei from spare embryos were tested using the same probe mixture as recommended for calculation the efficiency of hybridization; these two probes mixtures were validated in two sequential hybridization rounds as will be used in the main experiment.

2.1.2.1. Human Lymphocytes Isolation and Fixation

Lymphocytes were isolated from fresh, heparinized peripheral blood samples as previously described (Harper et al., 1994). These peripheral blood samples were withdrawn form patients with normal karyotype. The heparinized blood samples were diluted with 1X Red Blood Cell

Lysis Buffer (RBCLB) in a ratio of 1:1 for 10 to 15 minutes incubation at room temperature;

10X of RBCLB was prepared by dissolving 89.9 g of NH4Cl, 10 g of KHCO3, and 2 ml of 0.5M

EDTA in 1L of ddH2O with adjusted pH 7.3 (Sigma-aldrich, Oakville, ON, Canada). After,

56 samples were centrifuged at 1500 rpm for 5 min at room temperature. The supernatant was discarded, and the pellets were re-suspended in 1X PBS. The re-suspended pellets were then centrifuged at 1500 rpm for 5 min at room temperature; this process was repeated for three times until the supernatant was clear. A pre-warmed 10 ml of 0.075 M KCl hypotonic solution at 37ºC was added to the pellet, and mixed gently using plastic disposable pipette for 10-15 times. Tubes were incubated at 37ºC for 30 min, and centrifuged at 1500 rpm for 10 min at room temperature.

A fresh fixative solution of 3 methanol: 11acetic acid was prepared; 10 ml of the fixative solution was added to the lymphocytes pellets, and mixed gently using plastic disposable pipette for 10-15 times. Samples were incubated for 5 min at room temperature, and then centrifuged at

5000 rpm for 5 min at room temperature. The supernatant was discarded, and a fresh fixative solution was added; this process was repeated for three times. In the last time, 1-1.5 ml of fresh fixative solution was added to the pellets to be stored at –80°C until FISH validation process is required.

Lymphocytes were then fixed on super-frost, clean, and cold glass slides at 4°C. Three to four drops of lymphocytes in fixative solution were dropped on cold slides from a distance of about 30 cm to allow a physical rupture of the cellular membrane. The slides were then checked under microscope. The best area on the slides were selected using diamond pen where the more central area with higher number of well-spread nuclei. The slides at this stage were ready for hybridization validation.

2.1.2.2. Spare Human Embryos Manipulation and Fixation

Chromosomally abnormal spare embryos were donated with written consents by patients underwent PGD treatment cycles; these embryos were fixed as previously described (Bielanska et al., 2002a). Each of these embryos were transferred to a droplet of acid Tyrode’s for 2 min

57 (Sigma-aldrich, Oakville, ON, Canada) to remove the zona pellucida (ZP). Zona-free embryos were fixed directly on poly-L-lysine-coated glass slides (0.2 PLL/ml) (Sigma-aldrich, Oakville,

ON, Canada) under an inverted microscope (Olympus CKX; Olympus Canada) using 0.01mol/L

HCl/Tween 20 spreading buffer (Fisher Scientific, Ottawa, ON, Canada /Sigma-aldrich,

Oakville, ON, Canada).

The number and location of fixed nuclei for each embryo was recorded. The slides were air dried for 10-15 min, washed in PBS and dehydrated with a serial of diluted ethanol 50%, 70%, and absolute ethanol 100% for 2 min each step followed by air drying for 5 to 10 min. The slides containing the fixed nuclei were treated using pepsin (0.1 mg/ml in 0.01 HCl) (Sigma-aldrich,

Oakville, ON, Canada) at 37°C for 10 min to allow the cytoplasmic debris digestion. After the treatment, the slides were rinsed directly in distilled water for several times to wash off the excessive pepsin. Then, the slides were subjected to a fixative solution (1% paraformaldehyde

(PFA) in PBS) (Sigma-aldrich, Oakville, ON, Canada) at 4°C for 5 min, followed by washing and dehydration in the serial of PBS and ethanol dilutions at room temperature for 2 min each step. The slides at this stage were ready for hybridization validation.

2.1.2.3. FISH Analysis and Signals Scoring

FISH Analysis

FISH analysis was performed as previously described (Zhang et al., 2010). In the first round, the probe mixture of MultiVysion PB probe for chromosomes 13, 16, 18, 21, and 22 were combined with the nuclei on the glass slide and sealed with the coverslip for co-denaturation at

78°C for 5 min using HYBrite™ (Vysis, Downers Grove, IL, USA). The slides then were carried out in a moist chamber for hybridization at 37°C for overnight.

58 After hybridization, the slides were washed using stringent buffer (0.7X Standard Saline

Citrate (SSC)-0.3% Tween20) (Sigma-aldrich, Oakville, ON, Canada) at 68°C for 3 min to remove any unbound probes, followed by rinsing in less stringent washing buffer (2X SSC-0.1%

Tween20) at room temperature for three times. The hybridized nuclei were mounted in Antifade solution (p-phenylenediamide dihydrochloride; Vector, Burlingame, CA, USA). The nuclei were viewed under 100X magnification using the fluorescence microscope (Olympus CK60; Olympus

Canada), and the singles were scored using appropriate filters (DAPI, FITC, RED specific filters, double filter FITC/RED, triple filter DAPI/FITC/RED). Images were captures using CCD camera and CytoVision software (Applied Imaging, Santa Clara, CA, USA) equipped with the fluorescence microscope.

After reading the first round signals, the slides were exposed for direct light for 10 min to allow singles depletion, followed by washing in 2XSSC solution for 5 to 10 min to let a gentle detached of the coverslips. The slides then were washed and dehydrated using the serial of PBS and diluted ethanol (50%, 70%, and 100%) at room temperature for 2 min each step.

For the second round of hybridization, the probes mixture of chromosomes 15 and X were applied on the same nuclear material for co-denaturation at 78°C for 5 min. The slides then were carried out in a moist chamber for hybridization at 37°C for 3 h. The slides were washed using stringent buffer (0.4X SSC-0.3% Tween20) at 68°C for 2 min to remove any unbound probes, followed by rinsing in less stringent washing buffer (2X SSC-0.1% Tween20) at room temperature for three times. The hybridized nuclei were mounted in Antifade solution containing

0.25 ng/mL DAPI (4', 6-diamidino-2-phenylindole) (Sigma-aldrich, Oakville, Canada). The signals were viewed under the fluorescence microscope using appropriate filters. Images were captures using CCD camera and CytoVision software.

59 FISH Signals Scoring

FISH signals in lymphocytes and spare embryos nuclei were scored and interpreted as previously described (Bielanska et al., 2002a; Hopman et al., 1988; Scriven, Kirby, & Ogilvie,

2011). Only intact, undamaged nuclei were scored. Nuclei with dim signals for more than one of the chromosomes tested were considered as a hybridization failure and thereby were excluded.

Signals with diameter smaller than others in the same nucleus were considered non-specific probe binding and then were excluded. Two small, focal signals of the same chromosome, separated by a distance of less than one signal domain were considered as split one signal.

Lymphocytes that at least 95% of cells showed conclusive FISH signals were included in the study. Only 2-cell embryos which both cells showed conclusive FISH signals were included.

Embryos from 4-cells to blastocysts stage which at least 75% of cells showed conclusive FISH signals were included in the study.

Lymphocytes were considered normal when at least 95% of scored nuclei showed two

FISH signals for all screened chromosomes; the criteria for scoring FISH signals in lymphocytes has been determined previously described (Hopman et al., 1988). Embryos were classified according to the different chromosomal patterns detected in the hybridized nuclei: normal, abnormal, and mosaic; the criteria for scoring FISH signals in spare human embryos has been identified previously described (Bielanska et al., 2002a). Briefly, embryos were considered normal (diploid) when all or at least 75% of scored nuclei showed two FISH signals for all screened chromosomes.

Embryos were considered uniformly abnormal when all or at least 75% of scored nuclei showed an absence of two FISH signals for all screened chromosomes. Embryos were classified as haploid and polyploid when all or at least 75% of scored nuclei exhibit single, and three or

60 more FISH signals for all screened chromosomes. Embryos were classified as aneuploid when at least 75% of scored nuclei exhibit one or three FISH signals for one or two of screened chromosomes. Embryos were considered mosaic when a mixture of diploid and non-diploid of scored nuclei. Different patterns of mosaicism in human spare embryos have been determined previously (Bielanska et al., 2005; Bielanska et al., 2002a, 2002b).

2.2. Experimental Study Population (Examination Stage)

A total of 372 immature oocytes at either GV or MI stage and 120 unfertilized oocytes

(0PN) were donated with written consent by patients who underwent 130 controlled ovarian hyperstimulation (COH) treatment cycles between August 2012 and June 2014 at McGill

Reproductive Center. All abnormally fertilized oocytes (1PN/3PN) and frozen/thawed oocytes were excluded from the present project. Further morphological assessment and FISH analysis will be described in: 2.2.4: Oocytes Morphological Assessment; and 2.2.5:Oocytes Fixation and

FISH Analysis.

This study is ethically approved by the Research Ethics Board of McGill University Health

Center, Royal Victoria Hospital.

2.2.1. Patients Characteristics

Patients from any maternal age were included in the study and had more than 6 antral follicle counts (AFC). Any patients who had 6 AFC or less, and or underwent fertility preservation were excluded from the study. All treatment cycles were divided into two groups based on the maternal age at time of egg retrieval: cycles when women were younger than 36 yrs., and cycles when women were 36 yrs. and older with an average maternal age (31.65±3.04) and (38.81±2.01), respectively (Figure 2.1).

61 A total of 112 patients underwent 130 COH treatment cycles with an overall average of maternal age 35.6 4.2 years. Fourteen out of 112 couples were diagnosed with polycystic ovaries/polycystic ovary syndrome (PCO/PCOS), 22 with repeated pregnancy loss (RPL), 39 with male factor (MF), and the rest for other reasons including endometriosis and tubal factor

(Figure 2.2). Eighty-one out of 112 patients underwent only COH treatment cycles, whereas 31 out of 112 patients underwent PGD cycles (15 patients for single gene defects, 13 for translocation, and three for aneuploidy screening).

62

Figure 2.1: The percentiles of both maternal age groups per COH treatment cycles. The box plot demonstrates the distribution of maternal age in the two groups studied. The color coded divisions in the data represent the 50th percentile above and below the mean age of each group for this study; 31.81 years for the women younger than 36 years (young women), and 38.81 for the women 36 years and older (older women).

63

Figure 2.2: The infertility diagnosis among patients who underwent COH treatment cycles. The pie chart represents the infertility diagnoses among patients who had COH cycles, including both PGD and non- PGD patients; women diagnosed with polycystic ovaries/polycystic ovary syndrome (PCO/PCOS) were 12% of the total number of patietns included in this study; women with a history of repeated pregnancy loss (RPL) were 20% of the total number of patietns; women attended the fertility clinic due to male factor (MF) or due to other reasons such as endometriosis and tubal defect were about 35% and 33% of the total number of patients, respectively.

64 2.2.2. Controlled Ovarian Stimulation (COH) Treatment Cycles

All 130 COH treatment cycles were performed as previously described (Son et al., 2013).

Briefly, the basal transvaginal ultrasound scan was performed on the day 3 to 5 of the cycle for follicular count and endometrial thickness assessment. The scan was repeated on the day 7 to 9 for follicular count and endometrial thickness monitoring during undergoing ovarian stimulation drugs; the leading follicle should reach <12 mm in diameter and the endometrial thickness ≥6 mm. At this point 10,000 IU hCG was administered.

Eleven out of 130 treatment cycles were IVM cycles with a minimal ovarian stimulation among which two were without any stimulation. These IVM treatment cycles were performed as previously described (Shalom-Paz et al., 2011). The basal ultrasound was performed on day 2 to

3 of the cycle. The second scan was repeated on day 7 to 9 of the cycle. Any patient who showed a leading follicle reached <10 mm and or endometrium <6 mm were prescribed for 2 to 5 days of gonadotropins. Patients who showed dominant follicle >10 mm and or endometrium >6 mm continued without any stimulation. At this point 10,000 IU hCG was administered.

2.2.3. Oocytes Collection

The Oocyte retrieval was performed around 36 h post hCG priming as previously described

(Son et al., 2013). The oocytes were collected using transvaginal ultrasound–guided collection with a 19-gauge aspiration needle (K-OPS-7035-RWH-ET, Cook, Australia) under reduced aspiration pressure of 7.5 kPa. The aspirates were collected in tubes with pre-warmed heparinized saline. The aspirates containing cumulus-oocytes complexes (COCs) were washed with oocyte wash medium (Cooper Surgical, CT, USA) that contained HEPES buffer supplemented with recombinant human serum albumin, and then the COCs were isolated under a stereomicroscope for nuclear maturation assessment.

65 The nuclear maturity of the collected oocytes was performed under the dissecting microscope with high magnification of X80 using spreading method. In the absence of germinal vesicle of oocytes, the cumulus cells were removed with hyaluronidase using mechanical pipetting. These mature oocytes were inseminated on the same oocyte retrieval day (Day 0: 0–6 h). The un- denuded immature oocytes at (GV) and (MI) from cumulus cells were cultured in IVM medium

(Cooper Surgical, CT, USA) supplemented with 75 mIU/ml FSH and LH. Following culture on day 1 (24–30 h), the oocytes were denuded from cumulus cells with hyaluronidase, for further insemination. Oocyte that did not show any nuclear maturation were donated to the present project.

2.2.4. Oocytes Morphological Assessment For Chromosomal Complement Analysis

All immature oocytes were either at germinal vesicle (GV) or metaphase I (MI) stage as were assessed morphologically by the embryologists at the time of retrieval. Further morphological assessment at the time of receiving the oocytes (day1 and day3 post egg retrieval) was performed prior to and post oocyte fixation for chromosomal analysis (Figure 2.3). A total of four subsequent categories of oocytes were grouped based on the nuclear maturation and chromosomal configurations prior to and post oocyte fixation: (1) 105 at GV stage which were identified morphologically by the presence of GV; (2) 167 at the metaphase II (MII) stage (un- inseminated in vitro matured oocytes) which were identified morphologically by the presence of

1PBs; (3) 62 post germinal vesicle break down group A (post GVBD-A) stage which were identified morphologically by the absence of GV or 1PB with dispersed chromosomes; and (4)

38 post germinal vesicle break down group B (post GVBD-B) stage which were identified morphologically by the absence of GV or 1PB with undispersed, condensed chromosomes

66 (Figure 2.4). All unfertilized oocytes were fixed directly after the morphological assessment of the presence of its corresponding 1PBs (not shown in the diagram).

A total of 232 of successfully fixed and analyzed in vitro matured oocytes were divided into two groups based on the maternal age at the time of egg retrieval: women <36 yrs. (younger women) and women ≥36 yrs. (older women) with average maternal age (31.65±3.04) and

(38.81±2.01), respectively. In addition, a total of 67 out of 120 unfertilized oocytes that were successfully fixed and analyzed were divided into two the same maternal age groups (Figure 2.5)

(Table 2.1).

67

Total 372 immature oocytes (GV/MI) were donated from 130 COH treatment cycles at day 1 and/or day 3 post egg retrieval

The nuclear maturation of all donated oocytes were assessed morphologically prior and post fixation for FISH analysis

I. Morphological assessment of nuclear maturation prior to fixation under the stereomicroscope:

105 immature oocytes at 167 in vitro matured oocytes at 100 immature oocytes post Germinal Vesicle stage (GV) Metaphase II stage (MII) germinal vesicle break down (GVBD)

II. Morphological Assessment of Chromosomal configuration post fixation under the inverted microscope:

62 immature oocytes post germinal vesicle 38 immature oocytes post germinal vesicle break down group A (post GVBD-A) break down group B (post GVBD-B)

Figure 2.3: Schematic diagram of the morphological assessment of collected oocytes. The schematic diagram illustrates the method by which the oocytes in this study were assessed and identified morphologically prior to and post fixation (n=372). The initial morphological assessment (prior to fixation) yielded two groups: 105 immature oocytes at germinal vesicle (GV) stage were identified morphologically by the presence of GV, and 167 matured oocytes at the metaphase II (MII) stage were identified morphologically by the presence of the extruded 1PB. The remaining 100 oocytes were morphologically classified (post-fixation) as post-germinal vesicle breakdown (post-GVBD) which were identified morphologically by the absence of GV and 1PB; these oocytes were divided into two categories post fixation: post-GVBD-A represents those with dispersed condensed bivalents which were identified by the dispersed condensed bivalents post fixation, and post-GVBD-B were those with undispersed condensed bivalents which were identified by the undispersed condensed bivalents post fixation post fixation. Unfertilized oocytes that failed to fertilize after subjecting for sperms either by IVF or ICSI (n=120) (not shown in the diagram) were fixed directly after checking the corresponding 1PBs.

68

A.

B. B. B. 1PB

MII

C.

D. D.

Figure 2.4: Microscopic images of successfully fixed immature and mature oocytes. The light microscopic images from an inverted microscope display the released chromosomal material from oocytes at different maturation stage; A) immature oocyte at GV stage; B) mature oocyte with the corresponding first polar body (MII/1PB); and C) immature oocyte post GVBD-A, and D) immature oocyte post GVBD-B.

69 Table 2.1: The total number, mean, median and range of successfully fixed oocytes at different maturation stage per treatment cycles including unfertilized oocytes in relation to maternal. Total Number Mean Median Range Number of COH Treatment Cycles: Younger women1 65 N/A N/A N/A Older women2 65 N/A N/A N/A Maternal Age: Younger women1 N/A 31.65±3.04 32 (24-35.9) Older women2 N/A 38.81±2.01 38.3 (36-42.9) Number of Immature Oocytes per Treatment Cycle: Germinal Vesicle Oocytes (GV) Younger women1 62 2.3±2.02 1 (1-8) Older women2 30 1.50±0.83 1 (1-4) Post Germinal Vesicle Break Down Oocytes Group A (post GVBD-A) Younger women1 38 2.11±1.32 1.5 (1-5) Older women2 15 1.67±0.71 2 (1-3) Post Germinal Vesicle Break Down Oocytes Group B (post GVBD-B) Younger women1 16 1.45±1.04 1 (1-4) Older women2 21 1.31±0.60 1 (1-3) Number of Mature Oocytes per Treatment Cycle: Metaphase II Oocytes with the corresponding first Polar bodies (MII/1PBs) Younger women1 63 2.17±1.10 2 (1-4) Older women2 77 2.66±1.76 2 (1-7) Unfertilized oocytes with corresponding first Polar bodies (0PN/1PBs) Younger women1 36 1.9±1.5 1 (1-7) Older women2 31 1.8±1.5 2 (1-7) 1 Younger women indicates all women younger than 36 years at the time of egg retrieval. 2 Older women are all women 36 years or older at the time of egg retrieval.

70

Figure 2.5: Graphical representation of total successfully fixed oocytes at different maturation stages including unfertilized oocytes per treatment cycles. The diagram displays the number of donated oocytes according to maturation stage for the total number of oocytes used in this study. A Chi-squared test shows a significant difference between the maternal age groups (p=0002). Oocytes at a more immature stage are more collected from younger women, whereas more mature oocyte were collected from the older age group. Statistically significant difference in donated oocytes at each maturation stage including unfertilized oocytes between both maternal age groups using Chi-squared test (P=0.003).

71 2.2.5. Oocytes Fixation for FISH Analysis

All oocytes were fixed for FISH analysis as described previously (Dyban et al., 1996;

Magli et al., 2012; Munne, Dailey, Sultan, Grifo, & Cohen, 1995) In brief, each oocyte was washed for a couple of times using three drops of PBS containing 4 mg/ml Bovine Serum

Albumin (BSA) (Sigma-aldrich, Oakville, ON, Canada) to remove the residual oocyte wash medium and any attached cumulus cells (CCs). Each washed oocyte was exposed for 2 min using acid Tyrode’s solution (Sigma-aldrich, Oakville, ON, Canada) to allow a partial Zona Pellucida digestion. Afterwards, each Zona-free oocyte was transferred on Poly-L-Lysine-coated glass slide (0.2 PLL/ml) (Sigma-aldrich, Oakville, ON, Canada) under an inverted microscope

(Olympus CKX; Olympus Canada) using minimal volume of PBS. A maximum of four oocytes per slid were fixed separately using hypotonic spreading buffer (0.01mol/L HCl/Tween 20). The fixed nuclear material of each oocyte was recorded using diamond pen.

The slides were air-dried for 5 to 10 min, washed using PBS for 4 min and dehydrated using a serial of diluted ethanol 50%, 70%, and absolute ethanol 100% for 4 min each step followed by air drying for 5 to 10 min. FISH analysis then was performed as previously described in 2.1.2.3: FISH Analysis in lymphocytes and spare embryos.

2.2.6. Oocytes FISH Signals Scoring

Only nuclei of immature and mature oocytes, including unfertilized oocytes with round to elliptical in shape and regular in intensity FISH signals were scored (Dyban et al., 1996; Rowsey et al., 2013). Nuclei with dim or fused signals were considered as a hybridization failure and were not scored. Nuclei with irregular signals in intensity were considered as a non-specific hybridization and were not scored.

72 As previously described (Dyban et al., 1996), mature oocytes, including unfertilized oocytes were classified according to the FISH signal patterns detected into: (1) euploid when an oocyte and its corresponding first polar body showed two distinct signals of all chromosome analyzed; (2) aneuploid when an oocyte and or its corresponding first polar body exhibit one or three distinct signals of any the chromosomes analyzed; (3) haploid when an oocyte and or its corresponding first polar body displayed one distinct signals of all chromosome analyzed; (4) polyploid when an oocyte and or its corresponding first polar body showed three or more distinct signals of all chromosome analyzed.

Since there is no study showed the criteria of FISH signals scoring in immature oocytes, different patterns of FISH signals in these oocytes at different maturation stage will be presented in the results chapter.

2.3. Statistical Analysis and Data Interpretation (Post-examination Process)

Chi-squared test and Fisher’s exact test were performed where it is appropriate to calculate the differences between categorical variables. Results were expressed the mean values when the confidence interval equal 95%, and all two-tailed P-values were considered statistically significant when P <0.05.

73

CHAPTER THREE:

RESULTS

74 3.1. FISH Technique Validation in Human Lymphocytes and Spare Embryos, and FISH

Signals Scoring in Human Immature and Mature Oocytes

Due to the limited number of human oocytes donated to research, the methodology chosen for the present project had to be efficient to ensure that the experimental findings are reproducible and inconclusive results are eliminated. Therefore, the first phase of the project was aimed to optimize the Fluorescent In Situ Hybridization (FISH) technique for those chromosomes most frequently involved in pregnancy loss in interphase nuclei derived from human lymphocytes. Those derived from patients with a normal karyotype were used as negative controls. Our samples also included donated spare embryos. As a positive control, we used spare embryos that were donated after PGD treatment cycles and known to have abnormal chromosomal complements.

3.1.1. FISH Analysis Efficiency in Human Lymphocytes and Spare Embryos

The efficiency of FISH was tested on interphase nuclei from patients’ lymphocytes that were known normal karyotype. Specific FISH signals for chromosomes 13, 15, 16, 18, 21, 22, and X were detected in six normal control lymphocytes samples (two male and four female)

(Figure 3.1-A). The overall FISH efficiency rate was 97.4%. The efficiency of FISH was also tested on interphase nuclei from PGD patients’ spare embryos of known abnormal karyotype.

Specific signals for all analyzed chromosomes were detected in 74 abnormal control spare human embryos. (Figure 3.1-B). The overall FISH efficiency rate was 92.7%.

3.1.2. FISH Analysis Efficiency in Human Immature and Mature Oocytes

The efficiency rate of FISH was calculated in immature and mature oocytes (Table 3.1).

75

Interphase nuclei of of nuclei Interphase lymphocyte human

A.

Interphase nuclei of of nuclei Interphase preimplantation human embryo

B.

Figure 3.1: Expected normal FISH signals for chromosomes (13, 15, 16, 18, 21, 22, and X) from interphase nuclei from human lymphocytes and spare embryos. FISH images taken by florescence microscopy display the singles of chromosomes 13, 15, 16, 18, 21, 22, and X in oocytes at different maturation stage; signals for chromosome 17 were not included in the present data. Image A shows an euploid interphase nuclei from normal female at which two signals are shown uniformly for all chromosome 15 (green); chromosome 17 (red), and X chromosome (orange); and Image B shows an aneuploid interphase nuclei from cleavage stage embryos at which signals are shown for all chromosome 13 (red), chromosome 16 (aqua), chromosome 18 (blue), chromosome 21 (green), and chromosome 22 (gold), indicating monosomy for chromosome 13 and 16 (left image), and monosomy for chromosome 21 (right image).

76 Table 3.1: Total number of collected, fixed, and successfully hybridized using FISH analysis for all chromosomes. GV Post GVBD-A Post GVBD-B MII-1PB 0PN-1PB oocytes oocytes oocytes oocytes oocytes Total No. of Collected Oocytes 105 62 38 167 120 Total No. of Successfully Fixed Oocytes (%) 98 (93.3) 55 (88.7) 38 (100) 146 (87.4) 73 (60.8) Total No. of Successfully Analyzed Oocytes (%) 92 53 37 140 67 FISH Analysis Efficiency Rate % 93.9 96.4 97.4 95.9 91.7 FISH efficiency rate was calculated per oocytes group of maturation by dividing the total number of successfully analyzed oocytes by the total number of successfully fixed oocytes: immature oocytes at germinal vesicle (GV) stage, immature oocytes post germinal vesicle break down (GVBD-A) stage group A, immature oocytes post germinal vesicle break down (GVBD-B) stage group B, mature oocytes at metaphase II (MII) stage with the corresponding first polar bodies (1PB), and unfertilized oocytes (0PN) with the corresponding first polar bodies (1PB).

77 3.1.3. FISH Signals Scoring in Human Immature and Mature Oocytes

Oocytes were scored as chromosomally normal when all chromosomes tested showed the following pattern of FISH signals (Table 3.2): mature oocytes at MII stage with the corresponding first polar body - two signals (Figure 3.2-A-B), unfertilized oocytes with the corresponding first polar body - two signals (not shown) (Gutierrez-Mateo, Benet, et al., 2004;

Magli et al., 2012; Pujol et al., 2003), immature oocytes at the GV stage - two signals (Figure

3.2-C) (Roig, Robles, Garcia, Martinez-Flores, et al., 2005), immature oocytes post GVBD-A oocytes - four signals (Figure 3.2-D-E) (Garcia-Cruz, Casanovas, et al., 2010; Shin et al., 2010), and immature oocytes post GVBD-B oocytes - four signals (Figure 3.2-F) (Vlaisavljevic et al.,

2007). A total of 52 out of 140 (37.1%) mature oocytes at MII stage, 34 out of 67 (50.7%) unfertilized oocytes, 74 out of 92 (80.4%) immature oocytes at GV stage, 39 out of 53 (73.6%) immature oocytes post GVBD-A, and 27 out of 37 (72.9%) immature oocytes post GVBD-B showed the expected normal FISH signals (Figure 3.2-A-F).

Mature oocytes at MII stage were scored as chromosomally abnormal when an extra or missing FISH signal was observed, while the remaining signals are diploid (Figure 3.2-G). Out of the total mature and unfertilized oocytes, 62.9% and 49.3%, respectively showed different patterns of chromosomal abnormality (Table 3.3). About half of the aneuploid mature oocytes

(46%) and unfertilized oocytes (55%) displayed an extra or missing FISH signal for only one chromosome. Around 30% and 25% of the mature and unfertilized oocytes were aneuploid where only two chromosomes were involved, respectively. Almost a similar proportion of aneuploid mature oocytes and unfertilized oocytes were observed (21.6% vs. 22.2%, respectively) when more than two chromosomes were involved. Only two mature oocytes (2.3%) showed triploidy for all analyzed chromosomes.

78 On the other hand, immature oocytes at GV stage were scored as chromosomally abnormal when there was an extra or missing FISH signal, and immature oocytes post GVBD of both groups (A and B) were scored as chromosomally abnormal when there were two extra or missing

FISH signals (Table 3.2). Only 19.6 % (18/92) immature oocytes at GV stage, 26.4% (14/53) immature oocytes post GVBD-A, and 27% (10/37) immature oocytes post GVBD-B did not show the expected normal FISH signals (Figure 3.2-H) (Table 3.4). Half of the immature oocytes at GV stage showed an extra or missing FISH signal for only one chromosome. The other half of these oocytes exhibited a variation in FISH signals for all chromosomes analyzed. Moreover, all fourteen immature oocytes post GVBD-A showed a considerable variation in the FISH signals for all chromosomes screened, whereas all ten immature oocytes post GVBD-B exhibited an extra or missing FISH signal for only one chromosome (Table 3.4).

79 Table 3.2: The expected FISH signals pattern in chromosomally normal and abnormal immature and mature oocytes. Oocytes at Different Maturation Stages Expected Normal FISH Expected Abnormal Signals Patterns FISH Signals Patterns Mature Oocytes at MII stage 2 signals 1 or 3 signals1 Unfertilized Oocytes 0PN 2 signals 1 or 3 signals1 First Polar Body of MII and 0PN Oocytes 2 signals 1 or 3 signals1 Immature Oocytes at GV stage 2 signals 1 or 3 signals1 Immature Oocytes post GVBD-A stage 4 signals 2 or 6 signals2 Immature Oocytes post GVBD-B stage 4 signals 2 or 6 signals2 1 One signal should represent a monosomy, and three signals should reveal a trisomy in immature oocytes at GV stage and mature oocytes at MII stage, including unfertilized oocytes. 2 Two signals should represent a monosomy, and six signals should reveal a trisomy in immature oocytes post GVBD-A and B groups.

80

Mature oocytes oocytes Mature oocytes Mature

G. G.

at MII Stage and 1PB and Stage MII at

A. A. 1PB and Stage MII at

First BodyFirst Polar of

. .

B Oocytes Mature

At GV Stage GV At

C. Immature oocytes oocytes Immature C.

A StageA

-

A StageA StageA

- -

Immature oocytes oocytes Immature

Immature oocytes oocytes Immature

post GVBD post

D. D.

E. Immature oocytes oocytes Immature E. GVBD post GVBD post

H.

B Stage B

-

post GVBD post

F. Immature oocytes oocytes Immature F.

Figure 3.2: FISH signals in oocytes at different maturation stages. FISH images taken by florescence microscopy display the singles of chromosomes 13, 15, 16, 18, 21, 22, and X in oocytes at different maturation stage; signals for chromosome 17 were not included in the present data. Image A shows an euploid mature oocyte at MII stage at which two signals are shown uniformly for all chromosome 15 (green); chromosome 17 (red), and X chromosome (orange); Image B shows an euploid first polar body (1PB) at which two signals are shown uniformly for all chromosome 13 (red), chromosome 16 (aqua), chromosome 18 (blue), chromosome 21 (green), and chromosome 22 (gold); Image C shows an euploid immature oocytes at GV stage at which two signals are shown uniformly for all chromosome 15 (green), chromosome 17 (red), and X chromosome (orange); Image D shows an euploid immature at post-GVBD-A stage at which four signals are shown uniformly for all chromosome 15 (green); chromosome 17 (red) and X chromosome (orange); Image E shows an euploid immature post-GVDB-A stage at which four signals are shown uniformly for all chromosome 13 (red),

81 chromosome 16 (aqua), chromosome 18 (blue), chromosome 21 (green) and chromosome 22 (gold); Image F shows an euploid immature at post-GVBD-B stage at which four signals are shown uniformly for all chromosome 15 (green) and X chromosome (orange); Image G shows a trisomic mature oocytes at MII stage for chromosome 22 (gold; arrow) retrieved from younger woman and euploid for rest chromosomes at which two signals for chromosome 13 (red), chromosome 16 (aqua), chromosome 18 (blue), and chromosome 21 (green); and Image H shows immature oocytes post-GVDB-A stage with a variation in FISH signals number and shape for all chromosome 13 (red), chromosome 16 (aqua), chromosome 18 (blue), chromosome 21 (green) and chromosome 22 (gold).

82 Table 3.3: Different Patterns of Chromosomal Abnormality in Mature Oocytes. Different Patterns of The Total No. The Overall Rate of Chromosomal Abnormality in of Abnormal Mature Chromosomal Abnormality Mature Oocytes Oocytes % Pre Group of All Mature Mature Oocyte1 Oocytes2 Mature oocytes at MII stage Aneuploid One chromosome 41 46.6 33.9 Two chromosomes 26 29.5 21.5 Three or more chromosomes 19 21.6 15.7 Haploid 0 0 0 Polyploid 2 (Triploid) 2.3 1.7 Total 883 100 72.7 Unfertilized oocytes 0PN Aneuploid One chromosome 18 54.5 14.9 Two chromosomes 8 24.2 6.6 Three or more chromosomes 7 21.2 5.8 Haploid 0 0 0 Polyploid 0 0 0 Total 333 100 27.3 1 The percentage of overall rate was calculated by dividing the total number of aneuploid, haploid, or polyploid MII oocytes by the total number of abnormal mature oocytes at MII stage; similarly with unfertilized oocytes. 2 The percentage of overall rate was calculated by dividing the total number of aneuploid, haploid, or polyploid MII oocytes by the total number of abnormal oocytes (mature oocytes at MII stage and unfertilized oocytes); similarly with unfertilized oocytes. 3 No significance difference in the total number of chromosomally abnormal oocytes between mature oocytes at MII stage and unfertilized oocytes using Chi-squared test (P=0.06).

83 Table 3.4: FISH signals of all tested chromosomes in immature oocytes at all maturation stages assigned as non-chromosomally normal. Chromosomes involved in pre-meiotic errors Maternal Age group Immature Chr.13 Chr.15 Chr.16 Chr.18 Chr.21 Chr.22 Chr.X (sample #) oocytes Younger women 1 GV 2 2 2 2 1 2 2 2 GV 1 2 2 2 2 2 2 3 GV 2 1 2 2 2 2 2 4 GV 2 2 2 2 1 2 2 5 GV 2 2 1 2 2 2 2 6 GV 2 2 2 2 5 2 2 7 GV 4 2 2 2 4 4 2 8 GV 4 2 2 2 4 4 2 9 GV 4 4 2 2 4 4 4 10 GV 4 2 1 4 2 4 4 11* Post GVBD-A 4 4 4 3 4 4 4 12* Post GVBD-A 3 4 3 3 3 3 3 13 Post GVBD-A 4 4 2 2 4 4 4 14 Post GVBD-A 4 4 2 2 4 4 4 15 Post GVBD-A 4 3 4 3 3 4 4 16** Post GVBD-A 3 2 4 2 4 3 4 17 Post GVBD-A 4 2 2 1 2 3 3 18 Post GVBD-A 2 2 3 3 2 2 2 19 Post GVBD-B 4 6 4 4 4 4 4 20 Post GVBD-B 1 4 4 4 4 4 4 21 Post GVBD-B 4 4 4 4 5 4 4 Older women 1 GV 2 2 1 2 2 2 2 2 GV 2 2 1 2 2 2 2 3 GV 2 2 2 2 5 2 2 4 GV 3 2 3 2 2 2 2 5 GV 2 2 2 2 4 4 2 6 GV 2 4 2 2 2 2 4 7 GV 4 4 4 3 3 2 4 8 GV 4 2 1 2 4 4 4 9 Post GVBD-A 4 4 2 2 4 4 2 10 Post GVBD-A 4 2 2 2 4 4 2 11 Post GVBD-A 4 4 2 4 3 2 4 12 Post GVBD-A 2 2 2 2 1 2 2 13 Post GVBD-A 2 2 2 1 2 2 2 14 Post GVBD-B 4 4 4 1 4 4 4 15 Post GVBD-B 4 4 4 1 4 4 4 16 Post GVBD-B 4 4 4 4 1 4 4 17 Post GVBD-B 4 4 4 4 4 2 2 18 Post GVBD-B 4 4 4 4 6 4 4 19 Post GVBD-B 6 4 4 4 4 4 4 20 Post GVBD-B 4 3 4 4 6 4 4 *Samples from the same treatment cycle. **See (Figure 3.2-H).

84 3.2. FISH Signals Validation in Human Immature Oocytes

Because of the considerable variation in the FISH signals identified in immature oocytes versus mature oocytes (Table 3.4), the second objective of the present study was intended to statistically validate FISH signals for all chromosomes analyzed (13, 15, 16, 18, 21, 22 and X) in immature oocytes at different maturation stages: germinal vesicle (GV), post germinal vesicle break down group A (post GVBD-A), and post germinal vesicle break down group B (post

GVBD-B) by plotting histograms of FISH signals per tested chromosome in oocyte group, in relation to maternal age (Figure 3.3). Similar histograms were plotted for mature oocytes, including unfertilized oocytes groups as internal control groups (Figure 3.3). These histograms serve as diagnostic tools to identify sampling variation by illustrating the most frequently occurring event (i.e. modality) in a distribution. The significant difference between the two distributions of maternal age groups (young versus old women) was calculated using Chi- squared test (Table 3.5).

3.2.1. Distributions of FISH Signals in Mature Oocytes and Unfertilized Oocytes as Control Groups In mature oocytes at the MII stage and unfertilized oocytes (0PN) derived from both maternal age groups (younger and older women), the histograms of FISH signals for all chromosomes analyzed were all unimodal distributions (Figure 3.3-A-B). There was no significant difference between distributions of both maternal age groups per chromosome analyzed (Table 3.5). Two-FISH signals interval for all chromosomes analyzed represented the mode (the peak) of all unimodal distributions; such mode represents the most frequently observed FISH signals shown by mature oocytes at MII stage and unfertilized oocytes (0PN) derived from both maternal age groups. Therefore, both groups of oocytes were considered as control groups. Furthermore, all distributions showed a comparable degree of skew (the

85 distributions direction) and kurtosis (the distributions shape). These distributions of both maternal age groups were symmetrical around the mode. However, distributions of mature oocytes at MII stage and unfertilized oocytes (0PN) derived from younger women had more tendencies to cluster in the third interval (two-FISH signals), while distributions of oocytes of same groups collected from older women were more distributed among the four intervals (zero, one, three, and four-FISH signals).

The unimodal distributions represent one population of mature oocytes at MII stage and unfertilized oocytes derived from both maternal age groups. Approximately 37% (52/140) of total mature oocytes at MII stage and half of all unfertilized oocytes (34/67) showed uniformly two-FISH signals for all chromosomes analyzed. Sixty-three percent and 50% of total mature oocytes at MII stage and unfertilized oocytes, respectively displayed uniformly two-FISH signals for all chromosomes analyzed except for one (about 40% vs. 15%), two (21.5% vs. 6.6%), or more than two chromosomes (15.7% vs. 5.8%, respectively) that showed zero, one, three, or four-FISH signals. Only 1.7% of total mature oocytes at MII stage showed uniformly three-FISH signals. No significant difference was found between the total number of mature oocytes at MII stage and unfertilized oocytes that showed two-FISH signals and other than two-FISH signals

(Table 3.3) (Figure 3.3-A-B).

3.2.2. Distributions of FISH Signals in Immature Oocytes at Different Maturation Stages

Distributions of FISH Signals in Immature Oocytes at GV Stage

In immature oocytes at the GV stage retrieved from both maternal age groups, the distributions for all chromosomes screened were unimodal (Figure 3.3-C). There was a significant difference between both maternal age groups per chromosome analyzed (Table 3.5).

This statistical significance was due to a proportion of oocytes (about 9%; 8/92) with uniformly

86 one-FISH signal for all analyzed chromosomes. This pattern of FISH signals may be due to a misreading of the signals, more than a failure of hybridization where the chance to lose one signal per each chromosome is unlikely to occur. It is possible that these signals were too close to each other, causing difficulty in the assessment. When these oocytes were considered as euploid with two signals per chromosomes, no significant difference was found between both maternal age groups (Table 3.5). The mode of all unimodal distributions was in the two-FISH signals interval for all chromosomes analyzed. Also, all distributions were symmetrical around the mode, and had similar trends to cluster in the two-FISH signals interval.

The unimodal distributions represent one population of immature oocytes at GV stage derived from both maternal age groups. The majority of these oocytes (70%; 64/92) showed uniformly two FISH signals per chromosome screened (Figure 3.3-C). Only 2.2% (2/92) of all oocytes showed uniformly four FISH signals for all analyzed chromosomes. The remaining

19.6% (18/92) oocytes exhibit a variation of FISH signals: one, two, three, four, and five (Table

3.4). Out of these oocytes, nine and six showed uniformly two FISH signals for all chromosomes analyzed except one chromosome and two chromosomes, respectively. The remaining three oocytes showed a high variation in FISH signals.

Distributions of FISH Signals in Immature Oocytes post GVBD-A Stage

Alternatively, in immature oocytes post GVBD-A stage retrieved from both maternal age groups, the distributions of each screened chromosome was bimodal in both maternal age groups with no significant difference (Figure 3.3-D) (Table 3.5). The bimodal distribution represented two populations of immature oocytes post GVBD-A stage, where the two modes of the bimodal distributions were in the two- and four-FISH signals intervals for all chromosomes analyzed in both maternal age groups. Both modes were of varying prominence in oocytes derived from the

87 younger women’s group; the more prominent mode was in the four-FISH signals interval, while the less prominent mode was in the two-FISH signals interval. Both modes were of equal prominence in oocytes derived from the older women’s group.

The bimodal distributions showed a possibility of sampling variation; i.e., mixture of two oocytes populations. The first population of immature oocytes post GVBD-A stage showed uniformly four signals per chromosome screened, accounting for about 53% (28/53) of the total oocytes. The morphological characteristic of the chromosomes during fixation (Figure 2.4) suggests that these oocytes were arrested at the metaphase I (MI) stage, where the four-FISH signals represents the bivalents of all chromosomes analyzed (Figure 3.2-D-E). The second population of immature oocytes post GVBD-A stage showed uniformly two signals per tested chromosome, accounting for about 21% (11/53) of the total number of oocytes of the same group. This uniformity of FISH signals may represent oocytes arrested at the anaphase I (AI) stage, where the chromosomal material may belong to either the oocyte or its first polar body, which one of them assumed to have been lost during fixation. The remaining 26% (14/53) of the same group showed a considerable variation of FISH signals (Table 3.4).

Distributions of FISH Signals in Immature Oocytes post GVBD-B Stage

In the last group, the distributions for all chromosomes analyzed in immature oocytes post

GVBD-A stage retrieved from both maternal age groups were unimodal with no significant differences between both maternal age groups (Figure 3.3-E) (Table 3.5). The unimodal distributions portrayed a single population of immature oocytes post GVBD-B stage, where the mode was in the four-FISH signal interval for all chromosomes analyzed in both maternal age groups. All histograms were symmetrical around the mode, and had similar trends to cluster in the two-FISH signals interval.

88 This group of post GVBD-B immature oocytes has the smallest sample size of total immature oocytes (8.4%; 27/322). The morphological assessment of the nuclear maturation of all these twenty-seven oocytes prior to fixation showed an absence of first polar bodies and germinal vesicle. The chromosomal configuration in these oocytes post-fixation showed non- dispersed, condensed, metaphasic chromosomes (Figure 2.3-D) in contrast to the typical dispersed, condensed, metaphasic chromosomes following oocytes lysing. About 63% (17/27) out of the total of these oocytes showed uniformly four FISH signals per chromosome analyzed.

The remaining nine oocytes displayed uniformly four FISH signals except one chromosome with an extra or missing FISH signal.

89

Figure 3.3: Histograms of FISH signals in relation to maternal age per screened chromosome (13, 15, 16, 18, 21, 22, and X) in oocytes at all different maturation stages including unfertilized oocytes (the color coded histograms represents the FISH probes): Figure 3.3-A: Mature Oocytes at MII stage (control group): all histograms of FISH signals for all chromosomes analyzed in relation to maternal age show also unimodal distributions with similar degrees of skew and kurtosis. The X- axis represents the intervals of all possible FISH signals, and the Y-axis represents the frequency of mature oocytes at MI stage. As expected, two-FISH signal interval for all chromosomes analyzed representing the mode of all distributions of both maternal age groups. Therefore, this group of oocytes is considered as a control group. No significant difference between each two distributions of maternal age groups for all chromosomes analyzed in mature oocytes at MI stage (See Table 3.5 for p-values).

90

Figure 3.3-B: Unfertilized oocytes 0PN (control group): all histograms of FISH signals for all chromosomes analyzed in relation to maternal age show also unimodal distributions with similar degrees of skew and kurtosis. Two-FISH signal interval for all chromosomes analyzed is the mode of all distributions of both maternal age groups. Therefore, this group of oocytes is considered as a control group. No significant difference between each two distributions of maternal age groups for all chromosomes analyzed in unfertilized oocytes (0PN) (See Table 3.5 for p-values).

91

Figure 3.3-C: Immature oocytes at GV stage: all histograms of FISH signals for all chromosomes analyzed in relation to maternal age show also unimodal distributions with similar degrees of skew and kurtosis. Two-FISH signal interval for all chromosomes analyzed is the mode of all distributions of both maternal age groups. Significant difference was found between distributions of both maternal age groups for all chromosomes analyzed in immature oocytes at GV stage. No significant difference between each two distributions of maternal age groups for all chromosomes analyzed after the adjustment (See Table 3.5 for p-values).

92

Figure 3.3-D: Immature oocytes post GVBD-A: all histograms of FISH signals for all chromosomes analyzed in relation to maternal age show bimodal distributions. The modes were in two- and four-FISH signal intervals of all distributions of both maternal age groups. No significant difference between each two distributions of maternal age groups for all chromosomes analyzed in immature oocytes post GVBD-A stage (See Table 3.5 for p-values).

93

Figure 3.3-E: Immature oocytes post GVBD-B : all histograms of FISH signals for all chromosomes analyzed in relation to maternal age show also unimodal distributions with similar degrees of skew and kurtosis. Four-FISH signal interval for all chromosomes analyzed is the mode of all distributions of both maternal age groups. No significant difference between each two distributions of maternal age groups for all chromosomes analyzed in immature oocytes post GVBD-B (See Table 3.5 for p-values).

94 Table 3.5: The statistical difference between distributions of FISH signals in relation to maternal age per screened chromosomes in oocyte at different maturation stages, including unfertilized oocytes. P value Differences between Both Maternal Age Distributions for All Screened Chromosomes Using FISH (P-values1) Oocytes at Different Maturation Stage Chr. 13 Chr. 15 Chr. 16 Chr. 18 Chr. 21 Chr. 22 Chr. X Immature oocytes at GV stage 0.0254 0.0033 0.0125 0.0256 0.0115 0.0479 0.0040 Immature oocytes post GVBD-A stage 0.5493 0.6314 0.2182 0.7312 0.9098 0.3689 0.2037 Immature oocytes post GVBD-B stage 0.3541 0.3541 0.0000 0.5538 0.3001 0.2883 0.4709 Mature oocytes at MII stage 0.6724 0.4808 0.1677 0.9625 0.4905 0.2858 0.2003 Unfertilized 0PN oocytes 0.7108 0.4868 0.4173 0.2985 0.0741 0.6077 0.7108 Adjusted Immature oocytes at GV stage2 0.1138 0.1641 0.0966 0.1248 0.0500 0.1715 0.1632 1 The p-values were calculated using Chi-squared test when p < 0.05 with a 95% confidence interval; these values indicate whether a significant difference exists between two distributions in relation to maternal age (young and old women) per screened chromosome in each oocyte group. A significant difference represents a possibility of variation in FISH signals scoring, assuming these samples came from the same population. 2 The p-values were re-calculated using Chi-squared when the group of oocytes with one FISH signals for all chromosomes analyzed.

95 3.3. The Estimated Rate of Chromosomally Normal Immature and Mature Oocytes of

Total Study Group in Association to Maternal Age

Due to the possibility of misdiagnosis of the chromosomally abnormal immature oocytes using FISH analysis, the third objective of the present project was aimed to estimate the overall rate of the chromosomally normal immature oocytes, and then determine the association between the estimated incidence of the chromosomally normal immature and mature oocytes in relation to the maternal age.

3.3.1. Overall Estimated Rate of Chromosomally Normal Immature and Mature Oocytes

The overall rates of chromosomally normal immature oocytes at GV and MI stages was estimated to be 73% (72/92) and 53% (28/53), respectively. About 63% (17/27) of immature oocytes post GVBD-B group was chromosomally normal. Forty-two per cent (86/207) of all mature oocytes, including unfertilized oocytes were normal as were 50.7% (34/67) and 37.1%

(52/140) of all successfully analyzed mature oocytes at MII stage and unfertilized oocytes, respectively.

3.3.2. Overall Estimated Rate of Chromosomally Normal Immature and Mature Oocytes in

Relation to Maternal Age

Forty-two out of 62 (67.7%) and 22 out of 30 (73.3%) of immature oocytes at GV stage were normal in younger and older women, respectively. No significant difference was found between both maternal age groups. For immature oocytes post GVBD-A (MI as suggested),

57.9% (22/38) and 40% (6/15) were chromosomally normal in younger and older women, respectively, and no significant difference was also found between both maternal age groups.

About 81% (13/16) and 67% (14/21) in immature oocytes post GVBD-B showed four FISH signals for all chromosomes analyzed with no significant difference between both maternal age

96 groups (Table 3.6). On the other hand, 39 out of 63 (61.9%) and only 13 out 77 (16.9%) of mature oocytes at the MII stage were normal for all screened chromosomes in younger and older women, respectively, when two FISH signals were seen per tested chromosome in mature oocytes. There was a statistically significant difference between both maternal age groups (P <

0.0001) among mature oocytes that were analyzed. Similarly, twenty-four out of the total unfertilized oocytes (61.5%) derived from younger women and only ten unfertilized oocytes donated by older women were chromosomally normal, which showed a significant difference between both maternal age groups (P=0.037) (Table 3.6).

97 Table 3.6: The estimated rate of a normal chromosomal complement at different maturation stages in association to maternal age. Oocyte at Different Maturation Stages Expected FISH Signals Estimated Chromosomally P value1 for All Tested Normal Oocytes at Different Chromosomes Maturation Stages (%) Younger women Older women Immature oocytes at GV stage 2 signals 67.7 73.3 0.5842 Immature oocytes post GVBD-A stage 4 signals 57.9 40.0 0.2392 (Immature oocytes at MI stage)4 Immature oocytes post GVBD-B stage 4 signals 81.3 66.7 0.4612 Mature oocytes at MII stage 2 signals 61.9 16.9 < 0.00013 Unfertilized oocytes (0PN) 2 signals 61.5 35.7 0.0373 1 P value was calculated using Chai-squared test and Fisher's exact test where was appropriate. 2 No statistical difference in the rate of normal immature oocytes at all three different stages between both maternal age groups. 3 Statistically significant difference in the rate of normal mature oocytes between both maternal age groups. 4 Only immature oocytes at MI stage, as suggested, was calculated out of the total oocytes post GVBD-A group. The table displays the frequency of chromosomally normal oocytes in all oocytes analyzed at both immature and mature stages of development in both age groups. The difference between maternal age groups in immature oocytes is statistically insignificant. The only significant data in this set is the estimated rate of normal mature oocytes, including unfertilized oocytes, which show an increased percentage in the younger age group versus the older age group in the same category.

98 3.4. The Estimated Rate of Chromosomally Normal Immature and Mature Oocytes of

Sibling Group in Association to Maternal Age

To eliminate the possible effect of patient-to-patient variations on the overall rate of chromosomal complements in immature oocytes, the fourth objective was aimed to re-analyze the chromosomal complement in a cohort of only sibling immature and mature oocytes. Each patient donated a minimum of one immature oocyte and one mature oocyte in order to allow the estimation of rates of chromosomally normal immature and mature oocytes in association to maternal age.

3.4.1. Cohort of Sibling Immature and Mature Oocytes

A total of 163 successfully fixed and analyzed sibling immature and mature oocytes were divided into two groups based on the maternal age at the time of egg retrieval; only mature oocytes were included. Forty-nine immature oocytes at any maturation stage and 27 mature oocytes at MII stage were donated by the younger women’s group with an average maternal age of 32.4±3.0 yrs where each patient donated a minimum of one immature oocyte and one mature oocyte. Thirty-seven and 50 immature oocytes at any maturation stage and mature oocytes were derived from older women with an average maternal age of 38.9±1.8 yrs (Table 3.7). The total number of donated sibling immature and mature oocytes was significantly different between the two maternal age groups (p=0.003); younger women donated more immature oocytes at all maturations stages, whereas more mature oocytes were collected from the older age group

(Figure 3.4).

3.4.2. Overall Estimated Rate of Chromosomally Normal Immature and Mature Oocytes

The overall rate of chromosomally normal immature oocytes of all groups was estimated to be 76.6% (36/47); 53.3% (8/15), and 79.2% (19/24) of all successfully analyzed immature

99 oocytes at GV, post GVBD-A (MI oocytes as suggested), and post GVBD-B stages. Only 23.8%

(22/77) of all mature oocytes were chromosomally normal.

3.4.3. Overall Estimated Rate of Chromosomally Normal Immature and Mature Oocytes in

Relation to Maternal Age

The estimated rates of chromosomally normal immature oocytes at all maturation stages were found to be comparable when a minimum of one immature oocyte (at any maturation stage) and one mature oocyte at MII stage were donated per cycle (GV: 78.1% vs. 73.3%; post-GVBD-

A; 50% vs. 57.1%; post-GVBD-B; 88.9 vs. 73.3%, younger vs. older women). No significant differences were found between both maternal age groups. However, a statistically significant difference between both maternal age groups was found in the mature oocyte group (Table 3.8).

100 Table 3.7: Table of successfully fixed and analyzed oocytes at different maturation stages amongst sibling oocytes (when a minimum of one immature oocyte at any maturation stages and one mature oocyte were collected per treatment cycle). Total Number Mean Median Range Number of COH Treatment Cycles: Younger women1 26 N/A N/A N/A Older women2 20 N/A N/A N/A Maternal Age: Younger women1 N/A 32.4±3.0 32.2 (26.11-35.7) Older women2 N/A 38.9±1.8 38.2 (36.4-41.8) Number of Immature Oocytes per Treatment Cycle: Germinal Vesicle Oocytes (GV) Younger women1 32 2.9±2.8 1 (1-8) Older women2 15 1.7±1.0 1 (1-4) Post Germinal Vesicle Break Down Oocytes Group A (post GVBD-A) Younger women1 8 2.0±2.0 1 (1-5) Older women2 7 1.4±0.5 1 (1-2) Post Germinal Vesicle Break Down Oocytes Group B (post GVBD-B) Younger women1 9 2.3±1.5 2 (1-4) Older women2 15 1.4±0.8 1 (1-3) Number of Mature Oocytes per Treatment Cycle Metaphase II Oocytes with the corresponding 1st Polar bodies (MII/1PBs) Younger women1 27 2.1±1.0 2 (1-4) Older women2 50 2.9±2.0 2 (1-7) 1 Younger women indicates all women younger than 36 years at the time of egg retrieval. 2 Older women are all women 36 years or older at the time of egg retrieval. The table lists sibling oocytes (oocytes from the same cohort) according to age groups and oocyte maturation stages used in the current study. There were more young women with oocytes at the germinal vesicle (GV) and post-germinal vesicle break down A (GVBD-A) stages than the older cohort in the same category. The older women had more oocytes in the post germinal vesicle break down B (GVBD-B) and mature (metaphase II with corresponding first polar body) stages than the younger women in the same category.

101

Figure 3.4: Graphical representation of successfully fixed and analyzed sibling oocytes at all different maturation stages (when a minimum of one immature oocyte at any maturation stage and one mature oocyte were collected per treatment cycle). The diagram displays the number of donated oocytes according to maturation stage for the group of sibling oocytes used in this study. A Chi-squared test shows a significant difference between the maternal age groups (p=0.003). Oocytes in immature stage are more likely retrieved from younger women, whereas more mature stages of oocytes were collected more from the older age group. Statistically significant difference in donated oocytes at each maturation stage between both maternal age groups using Chi-squared test (P=0.003).

102 Table 3.8: The estimated rate of chromosomally normal immature oocytes across different maturation stages and mature oocytes in association to maternal age in sibling oocytes. Oocyte at Different Maturation Stages Expected FIISH Signals Estimated Chromosomally P value1 for All Tested Normal Oocytes at Different Chromosomes Maturation Stages (%) Younger women Older women Immature oocytes at GV stage 2 signals 78.1 73.3 0.4292 Immature oocytes post GVBD-A stage 4 signals 50.0 57.1 0.7852 Immature oocytes post GVBD-B stage 4 signals 88.9 73.3 0.3592 Mature oocytes at MII stage 2 signals 59.3 12.0 <0.00013 1 P value was calculated using Chai-squared test and Fisher's exact test where either was appropriate. 2 No statistical difference in the rate of normal immature oocytes at all three different stages between both maternal age groups. 3 Statistically significant difference in the rate of normal mature oocytes between both maternal age groups.

103 3.5. The Estimated Rate of Pre-meiotic Mitotic Errors and First Meiotic Errors in Mature

Oocytes and in Association to Maternal Age

The fifth objective of the present study was aimed to estimate the overall rate of pre- meiotic mitotic errors in mature oocytes, including unfertilized oocytes with the corresponding first polar bodies, and investigate the incidence of pre-meiotic mitotic errors in mature oocytes in association to maternal age. The first meiotic errors in mature oocytes with the corresponding first polar bodies were also analyzed.

3.5.1. Overall Estimated Rate of Pre-meiotic Mitotic Errors

The total number of chromosomal errors in mature oocytes, including unfertilized oocytes with the corresponding first polar bodies was 465 errors. These doublets cells (MII/0PN-1PB) showed a high coefficient of correlation (r = 0.81).

Overall Rate of Pre-meiotic Mitotic Errors in Relation to The Total Number of Chromosomal

Errors

Thirty-five (7.5%) out of the 465 errors were non-complementary errors, representing either pre-meiotic mitotic errors patterns in mature oocytes with the corresponding oocytes: a double gain of the screened chromosomes in one of the doublets cells (MII-1PB) (Four FISH signals in either cell) and the other cell was diploid of the same chromosome, or a gain of the screened chromosomes in both doublets cells (MII-1PB) (Three FISH signals per cell) (Figure

3.5). The incidence of the pre-meiotic mitotic errors in accordance with the first pattern was two times higher than errors displaying the second pattern of pre-meiotic mitotic errors (n=21 vs. n=7

(14), respectively) (Table 3.9).

104 Overall Rate of Pre-meiotic Mitotic Errors in Relation to The Total Number of Mature oocytes

Twelve percent (24/207) of the total mature oocytes, including unfertilized oocytes with the corresponding first polar bodies had pre-meiotic mitotic errors (Table 3.9).

Overall Rate of Pre-meiotic Mitotic Errors in Relation to The Patients Infertility

Seventeen out of 112 (13.9%) patients who underwent COH treatment cycles donated at least one mature oocyte carried pre-meiotic mitotic errors. Among all these patients, three donated more than one oocyte with pre-meiotic mitotic errors. Six mature oocytes were collected from only one patient (Table 3.9).

All 17 patients had a history with primary infertility, except four who attended the fertility clinic due to secondary infertility. Nine out of 17 patients with a history with RPL and only two out of 17 patients with a history with one miscarriage (MC) donated oocytes with pre-meiotic errors. Two out of these RPL female patients were translocation carriers, as presented in the

Table 3.9. Only two fertile patients attended the fertility clinic to donate and to undergo PGD treatment cycle for single gene defect (SGD) (Table 3.9).

Overall Rate of Pre-meiotic Mitotic Errors in Relation to The Maternal Age

The estimated rates of pre-meiotic mitotic errors were found to be comparable in mature oocytes, including unfertilized oocytes derived from younger and older women groups (9.1%;

9/99 vs. 13.4%; 15/108, respectively). No significant differences were found between both maternal age groups (P=0.281) (Table 3.9).

The Most Common Chromosomes Involved in Pre-meiotic Mitotic Errors

With the exception of the X chromosome, all autosomes analyzed were involved in pre- meiotic mitotic errors. The highest rate was found for chromosome 13, followed by 16, 18 and

22 as shown in Table (3.9).

105 Table 3.9: Pre-meiotic errors in mature oocytes and their corresponding first polar bodies. Maternal Age Infertility History Sample ID Mature Oocyte First Polar Body (yrs.) Younger Women1 29 5 yrs. of 1° infertility 105#2 Trisomy 21 Trisomy 21 30 5 yrs. of 1° infertility + RPL 49#1 2N Tetrasomy 18, 21 31 3.5 yrs. of 1° infertility 107#1 2N Tetrasomy 13 31 2 yrs. of 1° infertility + RPL + 46,XX, t (2; 17)2 56#1-1 Trisomy 22 Trisomy 22 31 1.5 yrs. of 1° infertility + MC 75#1 Trisomy 21 Trisomy 21 32 7 mon. of 1° infertility + RPL + 45,XX, t(13; 14) 36#1 Tetrasomy 13 2N 35 10 yrs. of 1° infertility 12#1 Tetrasomy 22 2N 35 2.5 yrs. of 1° infertility + RPL 51#6 Trisomy 16 Trisomy 16 35 1.5 yrs. of 1° infertility + MC + PCO 34#1-11 2N Tetrasomy 22 Older Women1 37 3.5 yrs. of 2° infertility + RPL 35#1-1 2N Tetrasomy 13, 18 35#2-2 Trisomy 13 Trisomy 13 35#2-8 Tetrasomy 16 2N 35#2-9 2N Tetrasomy 16 35#2-10 Trisomy 16 Trisomy 16 35#2-13 Tetrasomy 13 2N 37 Egg donor 13#1 Tetrasomy 16, 18 2N 13#4 Tetrasomy 18 2N 38 2 yrs. of 2° infertility + RPL 29#2 Tetrasomy 15 2N 38 3 yrs. of 2° infertility + RPL 18#1-1 Tetrasomy 18 2N 18#3-2 Tetrasomy 13 2N 40 2 yrs. of 2° infertility + RPL 42#1 Tetrasomy 13, 18 2N 43 2 yrs. of 1° infertility + RPL 23#1 2N Tetrasomy 223 39 SGD 32#6 Trisomy 16 Trisomy 16 41 2 yrs. of 1° infertility 76#8 Tetrasomy 13 2N 1 No significant difference in the overall rate of pre-meiotic mitotic errors between both maternal age groups using Chai- squared test (P=0.281). 2 No pre-meiotic mitotic or first meiotic errors for chromosome 17 were observed in immature and mature oocytes (data were not shown the present data). 3 See Figure (3.6).

106

Figure 3.5: Two patterns of FISH signals for pre-meiotic mitotic errors in mature oocytes with the corresponding first polar bodies. The first possibility of FISH signal pattern of mature oocyte with the corresponding first polar body carring pre- meiotic mitotic error when gain of an extra chromosoes in one of the doublets cells (MII-1PB) (Four FISH signals in either cell); here the oocyte is diploid for chromosome 22 while the corresponding first polar body is tetrasomic forchromosome 22. The second possibility is a gain of the screened chromosomes in both doublets cells (MII-1PB) (Three FISH signals per cell); here both oocyte and the corresponding first polar body are trisomic for chromosome 22.

107 MII oocytes (tetrasomy for chr.22) with the corresponding 1PB (diploid for chr.22)

Figure 3.6: FISH signals of pre-meiotic mitotic error in mature oocyte with thier corresponding first polar body. FISH image obtained by florescence microscopy displays the singles of chromosomes 13 (red), 21 (green), and 22 (gold) in mature oocytes with the corresponding first polar bodies. Here, the MII-1PB doublets display the first pattern of pre-meiotic mitotic errors (non-complementary error); however, the oocytes showed a tetrasomy for chromosome 22 and the corresponding first polar body is diploid for the same chromosome. The same oocytes reveals first meiotic error (complementary error); the oocyte exhibit a tetrasomy for chromosome 13 and the corresponding first polar body is nullisomy for the same chromosome.

108 3.5.2. Overall Estimated Rate of First Meiotic Errors

Further analysis was aimed to study the incidence in first meiotic errors in mature oocytes, including unfertilized oocytes in association to maternal age.

Four-hundred and thirty out of 465 chromosomal errors (92.5%) were found in 121 mature oocytes, including unfertilized oocytes (58.5%) with the corresponding first polar bodies, where chromosomally abnormal oocytes with these errors were associated with maternal age (Figure

3.7).

The incidence of the Mechanisms Leading to Complementary and Non- complementary First

Meiotic Errors

Premature division (PD) was the predominant mechanism leading to complementary first meiotic errors in both maternal age groups. A total of 30 and 100 complementary chromatid errors was found in younger and older maternal groups, respectively. However, non-disjunction

(ND) was the least common mechanisms leading to complementary first meiotic errors in the younger group compared to the older women. Anaphase Lag (AL) was the most common cause of non- complementary chromosome and or chromatid errors in either cells in both maternal age groups, where oocytes were more affected than the first polar bodies in the younger group (Table

3.10).

The incidence of chromosome and chromatid errors

There was a significantly higher rate of chromatid errors in comparesion to chromosomal errors in oocytes derived from the older and younger women, respectively (P=0.004) (Table

3.11).

109 The Most Common Chromosomes Involved in First Meiotic Errors

First meiotic errors for chromosome 13 were the highest in oocytes derived from both maternal age groups, followed by chromosome 21 and chromosome 16. The highest rate of first meiotic errors was found in chromosome 21, followed by 13, 15, and 16, as shown in Figure

(3.8).

110

Figure 3.7: The percentage of total chromosomally abnormal mature oocytes in relation to maternal age. The incidence of chromosomally abnormal mature oocytes is significantly associates with maternal age (See Table 3.6).

111 Table 3.10: The frequency of mechanisms leading to errors in mature oocytes in relation to maternal age. Total errors in younger women Total errors in older women Complementary errors PD1 30 per MII-1PB doublets 100 per MII-1PB doublets ND1 9 per MII-1PB doublets 6 per MII-1PB doublets Non- complementary errors AL leading to chromosome 16 in either cells (5 in oocytes + 11 in 1PBs) 3 in 1PBs errors AL leading to chromatid errors 8 in either cells (7 in oocytes + 1 in 1PBs) 8 in either cells (6 in oocytes + 2 in 1PBs) Pre-meiotic mitotic errors 10 (4 per MII-1PB doublets + 6 in either cells) 18 ( 3 per MII-1PB doublets + 15 in either cells) 1Significance difference in the total of first meiotic errors due to PD and ND between both maternal age groups using Fisher-exact test (P=0.005).

112 Table 3.11: The frequency of chromosomal and chromatid errors in association to maternal age. Chromosome errors Chromatid errors Loss Gain Total Loss Gain Total Oocytes derived from younger women 7 9 161 18 17 351 Oocytes derived from older women 0 14 141 53 47 1001 Total 7 23 30 71 64 153 1 Statistical significant difference in chromatid and chromosome errors between both maternal age groups using Fisher exact test (P=0.004). 2 Two tetraploid oocytes were excluded from the data.

113

Figure 3.8: First Meiotic errors among all analyzed chromosomes in mature oocytes between both maternal age groups. Chromosome 13 was the highest in oocytes derived from both maternal age groups, followed by chromosome 21 and chromosome 16. The highest rate of first meiotic errors was found in chromosome 21, followed by 13, 15, and 16.

114 The incidence of First Meiotic Errors in Association with In-Vitro Maturation

As described in the Material and Methods section of this thesis, immature oocytes at the time of egg retrieval were collected for chromosomal analysis on Day 1 and Day 3 post egg retrieval.

In vitro mature oocytes versus unfertilized oocytes derived from younger women

About 42% (15/36) of unfertilized oocytes collected from younger women were chromosomally abnormal, while 38.1% (24/63) of the total number of in vitro mature oocytes obtained from the same group were abnormal. No significant difference was found between both oocyte groups (P=0.44).

Day 1 in vitro matured oocytes versus. Day 3 in vitro matured oocytes derived from younger women

Thirty-two and 31 in vitro matured oocytes were collected for analysis from young women on Day 1 and Day 3 post egg retrieval, respectively. Ten (31.3%) in vitro matured oocytes collected at Day 1 were chromosomally abnormal, whereas 14 (45.2%) in vitro matured oocytes collected at Day 3 were deemed abnormal. No significant difference was observed between both oocyte groups (P=0.19).

115

CHAPTER FOUR:

DISCUSSION

116 Numerical chromosome abnormality is one of the leading factors that negatively affect human reproduction and significantly contribute to the high incidence of spontaneous abortions, stillbirths, and live births with congenital defects (Blank, 1966; Carr, 1971; T. J. Hassold &

Jacobs, 1984). Around 5% of these abnormalities are of paternal origin, while the vast majority

(95%) of numerical chromosome abnormalities are maternally linked (Antonarakis, 1991; T.

Hassold et al., 1984; P. A. Jacobs, 1992; Kupke & Muller, 1989; Magenis, Overton, Chamberlin,

Brady, & Lovrien, 1977).

These maternally related abnormalities are mainly due to meiotic errors occurring during oogenesis. Studies for over two decades have documented the prevalence of meiotic errors in human oocytes derived from women who underwent ART treatment cycles, and some of these studies are listed in Table 1.1. The clinical relevance of these errors to human reproduction and fertility has been also well reported. The direct association between these errors and advanced maternal age have been well documented (using different methodologies for chromosomal analysis) in human oocytes either donated for research or used for clinical practice (Angell,

1994; Dailey et al., 1996; Fragouli, Wells, Whalley, et al., 2006; Gabriel et al., 2011; Golbus,

1983; Handyside et al., 2012; T. Hassold & Chiu, 1985; Kuliev et al., 2011; Nakaoka, Okamoto,

Miharu, & Ohama, 1998; Pellestor et al., 2003; Pellestor & Sele, 1988; Roberts & O'Neill, 1995;

Zenzes, Wang, & Casper, 1992).

Some of these maternally related chromosome abnormalities might be attributed to errors occurring prior to meiosis. These errors arise as a result of mitotic errors occurring during pre- implantation embryonic development; some of these chromosomally abnormal cells may eventually become precursors to germ cells. Pre-meiotic mitotic errors occur during germ cell proliferation where any of these chromosomally abnormal cells may ultimately become gametes

117 (Figure 1.1). Case reports and studies for over four decades have reported the incidence of these pre-meiotic mitotic errors in peripheral blood, buccal, skin, and ovarian samples, as well as mature oocytes obtained from young women with a history of multiple trisomic conceptions, which have been recently reviewed by (Delhanty, 2011; Papavassiliou et al., 2015). However, limited studies have been conducted to estimate the incidence of pre-meiotic mitotic errors in foetal ovarian cells and mature oocytes derived from karyotypically normal women due to the difficulty in obtaining these normal samples (Table 1.2). In contrast to meiotic errors, the overall rate of pre-meiotic mitotic errors in the general population (with a normal karyotype), their contribution to the overall rate of chromosome abnormalities, the clinical relevance to human reproduction and fertility, and the association with maternal age remain unknown.

Therefore, the main aim of the present project is to investigate the incidence of pre-meiotic mitotic errors in human immature and mature oocytes derived from women who underwent controlled ovarian hyperstimulation (COH) treatment cycles in relation to maternal age. To our knowledge, the present study is the first to analyze the chromosomal complements using such a large sample size of human immature oocytes (at different maturation stages), and mature oocytes through FISH analysis of those chromosomes most frequently involved in pregnancy loss (13, 15, 16, 18, 21, 22 and X). There is only one previous study that analyzed the chromosomal copy number in immature oocytes at the GV or MI stage using the CGH technique

(Daina et al., 2014).

1.4. Pre-Experimental Stage: FISH Technique Standardization and Validation using

Human Lymphocytes and Spare Donated Embryos

Prior to the experimental stage, we aimed first to optimize the FISH technique using both interphase nuclei derived from human lymphocytes (as a negative control, which were derived

118 from patients with a normal karyotype), and donated spare embryos (as a positive control, which were donated after PGD treatment cycles known to be chromosomally abnormal). Since the human oocytes donated for research are highly valued, the methodology chosen for the present project should be efficient to ensure that any further experiments is reproducible, and that any inconclusive results were eliminated.

According to the guidelines of ESHRE PGD consortium best practice guidelines for fluorescence in situ hybridization-based PGD (Harton et al., 2011), FISH technique should be optimized for routine testing in a clinical cytogenetic laboratory. Although the present project is research-based at a large academic centre, there remains great difficulty in obtaining human oocytes donated for experimental purposes. Therefore, we optimized the FISH technique (to the standard required for clinical testing), and successfully analyzed over 600 lymphocyte interphase nuclei, and 1000 embryonic interphase nuclei with overall FISH efficiency rates of 97.4% and

92.7%, respectively. The lymphocyte interphase nuclei were obtained from karyotypically normal patients. These lymphocytes were considered as a negative control for the chromosome abnormalities under the optimal hybridization condition. Whereas, nuclei obtained from donated spare embryos were considered as a positive control for the chromosome abnormalities under the optimal FISH condition. These chromosomally abnormal embryos were donated by patients who underwent PGD treatment cycles for translocation or aneuploidy screening after the day of diagnosis; therefore, the clinical diagnosis of these embryos is known. It has also been recommended by the same ESHRE guidelines to use the commercially available probe set for at least chromosomes 13, 15, 16, 18, 21, 22, and X because these chromosomes represent those most associated with pregnancy loss with, and can be successfully detect up to a rate of 70% of the overall rate of chromosome abnormalities (Munne et al., 2010). The overall FISH efficiency

119 rate was acceptable (over 92% in both samples of lymphocytes and donated embryos), where the

FISH technique has been reported to error rate of approximately 7% under optimal conditions

(Munne et al., 2010). The quality of the donated embryos has been reported to have a considerable impact on the hybridization efficiency rate (Findikli et al., 2004; Wu et al., 2014).

Also, different methods of nucleus fixation have been shown to affect the overall FISH efficiency rate (Munne, Dailey, Finkelstein, & Weier, 1996; Velilla, Escudero, & Munne, 2002).

The variation in the quality of donated embryos, and the different nucleus fixation techniques (as described in the second chapter), explain the lower rate of FISH efficiency in nuclei derived from spare embryos as compared to nuclei obtained from lymphocytes.

4.2. Experimental Stage: Chromosomal Complement Analysis in Human Immature and

Mature Oocytes Using FISH Technique

The assessment of the chromosomal complement using FISH analysis depends crucially on several factors, including the hybridization efficiency being used for analysis, and the cell type being analyzed. In addition, the FISH signals scoring criteria is the most important factor that may affect the assessment of the chromosomal status. Misinterpreting FISH signals under optimal hybridization conditions affect the diagnostic accuracy rates, resulting in a high rate of false positive or false negative results (Munne et al., 2010). In our case, this can subsequently affect our estimations of the overall rate of pre-meiotic mitotic errors in immature oocytes

(Morris et al., 2012; Rowsey et al., 2013).

Since the present project is the first to investigate the incidence of pre-meiotic mitotic errors in human immature and mature oocytes derived from women who underwent COH treatment cycles (in relation to maternal age) using FISH analysis: (i) the FISH hybridization rate

120 in immature and mature oocytes should be efficient; and (ii) the FISH signals in immature and mature oocytes should be adequately scored and interpreted.

FISH Efficiency Rate in Immature and Mature Oocytes

After standardizing the FISH technique, we analyzed the chromosomal complements in a total of 182 immature oocytes at different maturation stages including the GV stage, post

GVBD-A stage, and post GVBD-B stage. Also, 207 mature oocytes were analyzed, including unfertilized oocytes with their corresponding first polar bodies (that showed a diploid pattern) using FISH for chromosomes 13, 15, 16, 18, 21, 22, and X. This yielded a FISH hybridization efficiency rate of over 92% in all groups of oocytes analyzed, with an approximate 7% error rate

(Table 3.1). These rates were consistent with previous studies conducted on similar populations of oocytes; this includes in vitro matured oocytes (Vlaisavljevic et al., 2007), unfertilized oocytes

(Martini et al., 2000; Pujol et al., 2003), and their corresponding first polar bodies (Magli et al.,

2006). This error rate has been reported to be acceptable in investigating the chromosomal complements using FISH analysis by previous investigators (Munne et al., 2010). We also encountered a considerable number of oocytes that were not analysable and compromised the overall efficiency the analysis (Table 3.1). Between 10% to 40% of immature oocytes in the post

GVBD-A stage, and mature oocytes in the MII stage (including unfertilized oocytes) were not analyzable. The chromosomes of all these oocytes were condensed and more dispersed, especially those that were derived from the older women cohort or collected at day three-post egg retrieval (aged oocytes)(Zhivkova et al., 2007). Comparable rates to our rates of fixation failure have been reported (Anahory et al., 2003; Martini, Flaherty, Swann, Payne, & Matthews,

1997; Vlaisavljevic et al., 2007).

121 FISH Signals Scoring and Interpretation in Immature and Mature Oocytes

After confirming the efficiency of FISH hybridization rate in experimental samples, we developed a FISH signal scoring criteria for immature oocytes at different maturation stages. To interpret these signals correctly, we used previously published immunofluorescence (IF) data gathered from experiments on human immature oocytes (Table 3.2), as our study is the first to study the chromosomal complement in human immature oocytes investigating the incidence of pre-meiotic mitotic errors.

Well-developed FISH signals scoring criteria and interpretation in mature oocytes

Scoring and interpreting FISH signals for analyzing mitotic errors (under optimal conditions in order to analyze the chromosomal complement) in human preimplantation embryos at the cleavage or blastocyst stages is simple and straightforward (Delhanty et al., 1993). Such criteria for FISH signal scoring and interpretation have been well established for over two decades (Delhanty, Harper, Ao, Handyside, & Winston, 1997).

However, scoring and interpreting FISH signals in human mature oocytes and their corresponding polar bodies (in order to investigate the incidence of meiotic errors using FISH) is relatively complex as compared to mitotic errors; though the criteria for scoring and interpretation these FISH signals have also been well developed for over a decade (Kuliev et al.,

2011; Verlinsky et al., 1996). This degree of complexity in analyzing meiotic errors as compared to mitotic errors is due to the reduction in chromosomal number (without any DNA replication) after first meiotic divisions thus preserving the diploidy status of the zygote (Kuliev &

Verlinsky, 2004). This gives rise to a mature oocyte at the MII stage and its corresponding 1PB - with two chromatids per chromosome (Magli et al., 2012). Each chromatid is round to elliptical, and this typically shaped FISH signal is representative of the normal mature oocytes at the MII

122 stage, and its corresponding first polar bodies (Figure 3.2-A-B) (Table 3.2). We found chromosomal abnormalities in a total of 52 out of 140 (37.1%) mature oocytes at MII stage, and

34 out of 67 (50.7%) unfertilized oocytes, indicating that a substantial number of oocytes (57%) were chromosomally abnormal for all chromosomes analyzed. These abnormal oocytes showed an extra or missing FISH signal in any of the chromosomes analyzed (Figure 3.2-G) (Table 3.2).

Our overall rate of chromosomally abnormal oocytes was consistent with previous studies that focused on in vitro matured oocytes and unfertilized oocytes (derived from a comparable range of maternal aged subjects), and who also used a similar probe panel of FISH analysis

(Magli et al., 2006; Pujol et al., 2003; Vlaisavljevic et al., 2007). On average, about half of the mature oocytes are chromosomally abnormal and the range of these oocytes shifted with the maternal age (Kuliev et al., 2011). However, other factors may contribute to the heterogeneity of the overall rates of chromosome abnormality (in human mature oocytes) as found in the literature, and with particular respect to maternal age (Table 1.1). One of these factors is the number of chromosomes screened. Our rate of chromosomally abnormal oocytes is significantly higher than the reported rates (5.3% vs. 9.6%) when only three chromosomes (18, 21, and X)

(Honda et al., 2002) and four chromosomes (1, 5, 19, and X) (Zhivkova, 2003) were analyzed, respectively. Relatively comparable rates were found between the present study and previously reported rates when six or more chromosomes were screened; 48% and 44% of tested mature oocytes were chromosomally abnormal when a panel of six FISH probes (for chromosomes 1, 7,

13, 18, 21, and X) (Martini et al., 2000), and a panel of nine FISH probes (for chromosomes 1,

13, 15, 16, 17, 18, 21, 22, and X) (Pujol et al., 2003) were analyzed. However, our overall rate of chromosomally abnormal oocytes is relatively lower than the reported rates of about 67%

(Gabriel et al., 2011) 70% (Fragouli, Alfarawati, Goodall, et al., 2011), and 76% (Handyside et

123 al., 2012) when all 23 sets of chromosomes are screened using aCGH in subjects comparable in maternal age to our study.

However, meiotic errors are chromosome-specific in contrast to post-zygotic mitotic errors; each chromosome acts in a unique manner during meiotic divisions (Nagaoka et al.,

2012). The chromosome size, the centromere location, and the CG-rich regions have been suggested to play a role in the number of chiasmata and density of cohesions; however, the specific underlying molecular basis of this unique characteristic remains unclear (Jones & Lane,

2013). It has been demonstrated that a panel of five FISH probes for chromosomes (13, 16, 18,

21, and 22), and a panel of nine FISH probes for chromosomes (13, 15, 16, 17, 18, 21, 22, X and

Y) can detect about 70% and 73% of chromosome abnormalities, while 100% can be detected by aCGH (Munne et al., 2010). Therefore, including chromosomes 13, 16, 18, 21, 22, X and Y in

FISH analysis for chromosomes screening is recommended by the guidelines of ESHRE PGD consortium best practice guidelines for fluorescence in situ hybridization-based PGD (Harton et al., 2011). This, in part, explains the relatively lower overall rate of chromosome abnormalities in human oocytes (35%) when a panel of five FISH probes for chromosomes (1, 7, 15, X, and Y)

(Martini et al., 1997) is analyzed, as compared to our findings. Comparable rates have been reported to our 57% chromosomal abnormality rate, versus 47% and 53%,when two different five-panel FISH probes ((for chromosomes 13, 16, 18, 21, and 22) (Kuliev et al., 2011), and chromosomes (13, 16, 18, 21, 22, and X) (Montag et al., 2005) ), were analyzed.

In mature oocytes, we were not only able to diagnose, but could also classify the patterns of chromosomal abnormalities (Table 3.3). As expected, aneuploidy (the most common human chromosomal abnormality) was the most prevalent anomaly in mature and unfertilized oocytes

(47%). Out of these aneuploid oocytes, about half showed an extra or missing FISH signal for

124 only one chromosome, 30% showed an extra or missing FISH signal for two chromosomes, and

22% showed an extra or missing FISH signal for more than two chromosomes. Only two mature oocytes (2.3%) were triploid for all analyzed chromosomes. These results support the majority of previously published data on this topic (Table 1.1), as well as those conducted on pre- implantation embryos using an SNP array (Forman et al., 2013; Mantikou et al., 2012).

Developing FISH Signals Scoring Criteria in Chromosomally Normal Immature Oocytes

Immature oocytes were scored as chromosomally normal when all chromosomes tested showed the following pattern of FISH signals: immature oocytes at the GV stage - two signals

(Figure 3.2-C) (Roig, Robles, Garcia, Martinez-Flores, et al., 2005), immature oocytes at the post

GVBD-A stage - four signals (Figure 3.2-D-E) (Garcia-Cruz, Brieno, et al., 2010; Garcia-Cruz,

Casanovas, et al., 2010; Shin et al., 2010), and immature oocytes at the post GVBD-B stage - four signals (Figure 3.2-F) (Vlaisavljevic et al., 2007).

We observed normal FISH signals for all chromosomes analyzed in a total of 74 out of 92

(80.4%) immature oocytes at the GV stage, 39 out of 53 (73.6%) immature oocytes at the post

GVBD-A stage, and 27 out of 37 (72.9%) immature oocytes at the post GVBD-B stage (Figure

3.2-A-F). Although each bivalent is maintained by the chiasmata, and centromeric and telomeric cohesions, tow signals are expected in chromosomally normal immature oocytes at the GV stage, where each signal represents a univalent of the condensed bivalent confined within the GV. On the other hand, four signals are expected in chromosomally normal immature oocytes classified as post GVBD-A and post GVBD-B, due to the absence of the GV and 1PB, where each signal represents a chromatid of the bivalent (Garcia-Cruz, Brieno, et al., 2010; Garcia-Cruz,

Casanovas, et al., 2010; Shin et al., 2010).

125 Difficulty of scoring FISH signals in chromosomally abnormal immature oocytes

However, scoring the chromosomally abnormal immature oocytes using FISH analysis (in order to investigate the incidence of pre-meiotic mitotic errors) is very complex and not as straightforward as scoring mature oocytes or preimplantation embryos (Morris et al., 2012;

Rowsey et al., 2013). About 20% (18/92) of immature oocytes at the GV stage, 26.4% (14/53) of immature oocytes at the post GVBD-A stage, and 27% (10/37) of immature oocytes at the post

GVBD-B stage, did not show the expected normal FISH signals (Figure 3.2-H) (Table 3.4). Half of the immature oocytes at GV stage showed an extra or missing FISH signal for only one chromosome. However, the other half of these oocytes exhibited a considerable variation in

FISH signals for all the chromosomes analyzed.

The difficulty in scoring and interpreting FISH signals under optimal conditions in immature oocytes is mainly due to the variation in the chromosomal behaviour during the transition phase from the dictyate stage of prophase I (in GV oocytes), to metaphase II in MII oocytes, as shown in mouse (Arnault, Doussau, Pesty, Lefevre, & Courtot, 2010; Bonnet-Garnier et al., 2012; X. Y. Liu et al., 2005; Qi et al., 2013) and human oocytes (Combelles, Cekleniak,

Racowsky, & Albertini, 2002; Garcia-Cruz, Brieno, et al., 2010). This transition phase between the onset of the GVBD and the first polar body extrusion is about 36 hours in duration in human oocytes. The chromosomes behave differently during various stages of maturation, and the chromosomal material has been shown to behave differently during the GV stage and as it progresses to the onset of GVBD stage. This variation in the chromosomal behaviour may affect the shape and intensity of FISH signals (Robles et al., 2007; Roig, Robles, Garcia, Martin, et al.,

2005; Rowsey et al., 2013). For example, we observed in immature oocytes - non-round, non- elliptical, irregularly shaped FISH signals that were hard to assess (Figure 3.2-E, 3.2-H) (Table

126 3.4), as compared to round, regularly shaped FISH signals in mature oocytes at the MII stage

(Figure 3.2-A, 3.2-G).

In addition, the difficulties in assessing the chromosomal complements in immature oocytes depend on whether the hybridized probes co-localize (especially sub-telomeric probes) with the chiasmata (Robles et al., 2007; Roig, Robles, Garcia, Martin, et al., 2005). This may affect the number of true FISH signals. For instance, we observed animmature oocyte at the post

GVBD-A stage which had normal signals for most chromosomes, and one that showed three signals instead of four (Figure 3.2-H) (Table 3.4). One possibility to interpret these signals is that two signals may represent two achiasmatic non-sister chromatids, and one signal represents two chiasmatic non-sister chromatids (False positive) (Rowsey et al., 2013). Another possibility is that all three signals represent a univalent of a true trisomic immature oocyte (Garcia-Cruz,

Casanovas, et al., 2010).

Depleted cohesions may play a role in the variation of FISH signals (Garcia-Cruz, Brieno, et al., 2010). It has been well demonstrated that the premature separation of sister chromatids of either of the homologous chromosomes is linked to the depletion of cohesions concentrated around the centromeres of the homologous chromosomes in aged immature oocytes (Chiang et al., 2010; Garcia-Cruz, Brieno, et al., 2010; Hodges et al., 2005; Lister et al., 2010; L. Liu &

Keefe, 2008; Tachibana-Konwalski et al., 2010; Tsutsumi et al., 2014). This can be misleading in the interpolation of the three FISH signals of diploid immature oocytes at the GV stage with a premature separation of sister chromatids of either of the homologous chromosomes with three

FISH signals of true trisomic immature oocytes at the GV stage (Table 3.4), especially those were derived from older women. All of these factors contribute to the difficulty in scoring and interpreting FISH signals under optimal conditions in immature oocytes, resulting in over and/or

127 under-estimation of the overall rate of pre-meiotic mitotic errors in these cells at different maturation stages.

Misinterpretation of FISH signals has been is evident in a study by (Hulten et al., 2008), where they performed dual-probe FISH in foetal ovarian cells derived from karyotypically normal foetuses, for chromosome 21. Hulten’s group reported a fairly high rate of trisomic cells between the range of 0.2 to 0.9%. The same group proposed, at a later time, the OMSM model in order to explain the maternal age effect on the increasing rate of chromosome abnormalities in humans (Hulten et al., 2010). The group suggested that the trisomic pre-meiotic cells can initiate prophase I; these trisomic meiotic oocytes may be able to reach the dictyate stage and escape the atresia around the time of the birth. They justified the maternal age effect by suggesting a selection might occur against the trisomic immature oocytes for maturation (Figure 1.6) for further ovulation, resulting in an increase of these trisomic oocytes in the ovarian reserve of women with advance maternal age compared to younger women. Only two studies have been carried out to re-examine the frequency of occurrence of trisomic ovarian cells derived from foetuses with a normal karyotype, using FISH for chromosomes 16, 21, and 22 (Morris et al.,

2012) and FISH probes for chromosomes 13, 16, and 21 (Rowsey et al., 2013). Both studies have failed to replicate the high rate of trisomy in the foetal ovarian cells, suggesting that gonadal mosaicism does not explain the maternal age effect on increased rate of aneuploidy in humans.

Previously published reports have also demonstrated the comparable low rates of foetal oocytes at the leptotene stage with trisomy 18 only (0.7%; 4/ 693) and derived from foetuses with normal karyotypes using FISH probes for chromosome13 and 18 (Roig, Robles, Garcia, Martin, et al.,

2005) with that of foetal oocytes at the leptotene stage with trisomy 21 only (0.2%; 1/425) using

FISH probes for chromosome 21 and X (Robles et al., 2007). These conflicting results mey be

128 explained by the possibility of a mixed population of somatic, pre-meiotic, and meiotic ovarian cells since the gestational age of the foetuses of these cells ranged from 12 to 22 weeks (Figure

1.2). Given the variability in chromosomal organization in these cells, Hulten’s group scored and interpreted FISH signals equally (three FISH signals per trisomic cell). In fact, the same group

(Hulten et al., 2014) reported recently a consistent result with (Morris et al., 2012; Rowsey et al.,

2013); that is, a considerably low rate (0.07%) of trisomy in foetal ovarian cells derived from 12 foetuses between 9 to 11 weeks of gestation.

Taken together, these reports suggest that scoring and interpretation of FISH signals is a critical factor affecting the assessment of the chromosomal status, particularly so in immature oocytes. Misinterpreting these signals under optimal hybridization conditions can affect the diagnostic accuracy rates, resulting in a high rate of false positive or false negative results. In our case, this can subsequently affect the overall rate of pre-meiotic mitotic errors in immature oocytes. Hence, extra caution should be taken during the process of scoring and interpreting these signals.

FISH Signals Validation in Immature Oocytes

Because of the considerable variation in FISH signals in immature oocytes (Table 3.4), the second objective of the present study was aimed to further validate the FISH signals for all chromosomes analyzed (13, 15, 16, 18, 21, 22 and X) in immature oocytes at different maturation stages: germinal vesicle (GV), post germinal vesicle break down group A (post

GVBD-A), and post germinal vesicle break down group B (post GVBD-B) by plotting histograms of FISH signals per tested chromosome of each oocyte group in relation to maternal age (Figure 3.3-C-E). Similar distributions were plotted for mature oocytes, including unfertilized oocytes groups as a control (Figure 3.3-A-B). These histograms serve to identify

129 potential sampling variation by depicting the mode in a distribution. The differences between each group of oocytes, per chromosome analyzed in relation to maternal age were calculated using Chi-squared test.

Distributions of FISH signals in Mature Oocytes and Unfertilized Oocytes as Control Groups

In mature oocytes at the MII stage and unfertilized oocytes (0PN) derived from both younger and older women groups, the histograms of FISH signals for all chromosomes analyzed were unimodal distributions, with no significant differences between distributions of the two maternal age groups, per chromosome analyzed (Table 3.5). The two-FISH signals interval was the mode in the unimodal distributions for all chromosomes analyzed. This mode indicates that two-FISH signals (known status of diploidy) were the most frequently displayed by mature oocytes at the MII stage and unfertilized oocytes (0PN) derived from both maternal age groups.

Therefore, both groups of oocytes were considered as controls. In addition, all distributions showed a comparable degree of skew and kurtosis. Distributions of both maternal age groups were symmetrical around the mode, whereas distributions of mature oocytes at the MII stage and unfertilized oocytes (0PN) derived from younger women tended to cluster in the third interval

(two-FISH signals). Distributions of oocytes of same groups collected from older women were more spread among the four intervals (zero, one, three, and four-FISH signals). This is explained by the direct association between the increased number of chromosome abnormalities and maternal age. More mature oocytes with two-FISH signals were obtained from the younger group while more mature oocytes with other than two-FISH signals were obtained from the older women(Kuliev et al., 2011).

130 Distribution of FISH Signals in Immature Oocytes at the Germinal Vesicle Stage

The distribution of FISH signals for all chromosomes screened in immature oocytes at the

GV stage retrieved from both maternal age groups were unimodal in nature. There was a significant difference between both maternal age groups per chromosome analyzed. This statistical significance can be explained by the presence of a small population of oocytes (9%;

8/92), which exhibited uniformly one-FISH signal for all analyzed chromosomes. This pattern of

FISH signals may be due to misreading of the signals, more than a failure of hybridization where the chance to lose one signal per each chromosome is unlikely to occur. It is possible that these signals were too close to each other, giving rise to one overlapped signal per screened chromosome that was difficult to be assessed. When these oocytes were considered as euploid with two signals per chromosomes, no significant difference was found between both the maternal age groups.

Similar to the control group, the unimodal distributions represent one population of immature oocytes at the GV stage derived from both maternal age groups. The mode of all unimodal distributions was in the two-FISH signals interval for all chromosomes analyzed.

Moreover, all distributions were symmetrical around the mode, and had similar trends to cluster in the two-FISH signals interval. All these suggest that immature oocytes at the GV stage derived from both maternal age groups are comparable in terms of FISH signals displayed. The majority of these oocytes (70%; 64/92) showed uniformly two-FISH signals per chromosome screened

(Figure 3.3-C), and only 2.2% (2/92) of all oocytes showed uniformly four FISH signals for all analyzed chromosomes. These oocytes may be at the late GV stage or the early GVBD stage where these four signals may represent each chromatid per bivalent (Garcia-Cruz, Brieno, et al.,

131 2010; Qi et al., 2013). The possibility of depleted cohesions for all chromosomes is unlikely, and has not yet been reported.

The remaining 19.6% (18/92) of oocytes showed a variation of FISH signals - one, two, three, four, and five (Table 3.4). Out of these oocytes, nine showed normal (2) FISH signals for the majority of analyzed chromosomes, and one chromosome showed only 1 FISH signal. Six oocytes showed two FISH signals for most chromosomes, except two chromosomes displayed only one FISH signal for those analyzed. One of the possibilities to explain the single FISH signal is that these GV oocytes are truly monosomic for the chromosome involved (the single

FISH signal represents one univalent; these pre-meiotic mitotic errors for similar chromosomes have been reported previously using CGH (Daina et al., 2014). Another possibility is that these oocytes are truly diploid, while one FISH signal overlapped the other signals, or the other signals failed to hybridize. The remaining oocytes showed a high variation in FISH signals. This considerable variation in FISH signals can be explained by the combination of previously mentioned factors limitations pertaining to FISH signal interpretation in immature oocytes.

These oocytes cannot bedefinitively diagnosed using FISH techniques only. Only one oocyte, which was retrieved from an older woman, showed three signals for chromosomes 13 and 16, and two FISH signals for the rest of the screened chromosomes. This single egg can be diagnosed as a trisomic GV oocyte, indicating an incidence of pre-meiotic mitotic error; however, these signals could be a false positive due to a probe co-localization with the chiasmata

(Rowsey et al., 2013). Therefore, investigating the incidence of chromosomally abnormal immature oocytes at the GV stage can be overestimated due to the difficulty of interpreting FISH signals attributed to pre-meiotic mitotic errors.

132 Distributions of FISH Signals in Immature Oocytes at the Post GVBD-A stage

The distributions of FISH signals for each screened chromosome in immature oocytes at the post GVBD-A stage retrieved from both maternal age groups displayed bimodal distributions with no significant difference (Figure 3.3-D) (Table 3.5). As both populations of oocytes derived from both maternal age groups are comparable, the bimodal distribution represents two mixed populations of immature oocytes at the post GVBD-A stage. This step of the FISH signals validation is significance to identify the sampling variation. These histograms displayed two modes (the two- and four-FISH signal intervals) for all chromosomes analyzed in both maternal age groups. In oocytes derived from younger women the more prominent mode was the four-

FISH signals interval, and the less prominent mode was the two-FISH signals interval. The modes per distribution, on the other hand, were of equal prominence in oocytes derived from the older women there were equal populations of oocytes displaying four FISH signals and two

FISH signals.

From the total studied oocytes, the first population of immature oocytes at the post GVBD-

A stage (from both maternal age groups) showed uniformly four FISH signals per chromosome screened, accountingfor 53% (28/53) of the total. The number of FISH signals and the morphological characteristics of the chromosomes during fixation (Figure 2.4) suggest that these oocytes were arrested at the metaphase I (MI) stage, and the four-FISH signals represent a chromatid per bivalent of all chromosomes analyzed (Figure 3.2-D-E) (Garcia-Cruz, Brieno, et al., 2010; Garcia-Cruz, Casanovas, et al., 2010; Shin et al., 2010).

The second population of immature oocytes at the post GVBD-A stage showed uniformly two signals per tested chromosome which accounts for about 21% (11/53) of the total oocytes.

This uniformity of FISH signals may represent oocytes arrested at the anaphase I (AI) stage,

133 where the chromosomal material may belong to either the oocyte or its first polar body (in which one of them is assumed to have been lost during fixation)(Garcia-Cruz, Brieno, et al., 2010; Qi et al., 2013). The remaining 26% (14/53) of the same group showed a considerable variation of

FISH signals (Table 3.4), where the combination of FISH signal interpretation factors may be responsible for the observed results. Again, FISH cannot be used exclusively to definitively diagnose these chromosomal errors.

As the first meiotic division takes place, oocytes with only two or six signals may represent a monosomic or trisomic immature oocyte at the MI stage. Therefore, estimating the incidence of the chromosomally abnormal immature oocytes at this stage can be overestimated due to the possible confusion with first meiotic errors, in conjunction with a possible high rate of false positives due to fixation and/or hybridization failures.

Distributions of FISH Signals in Immature Oocytes at the Post GVBD-B stage

In the last group, the distributions for all chromosomes analyzed of immature oocytes (post

GVBD-A stage) were unimodal with no significant difference (Figure 3.3-E) (Table 3.5). Similar to immature oocytes at the GV stage, the unimodal distributions portrayed a single population of immature oocytes at the post GVBD-B stage, where all samples displayed four-FISH signal intervals for all chromosomes analyzed. All histograms were symmetrical around the mode, and had similar trends and clusters in the four-FISH signals interval in both maternal age groups.

This implies that immature oocytes in the post GVBD-B stage, derived from both maternal age groups, are comparable in terms of displayed FISH signals.

This group of post GVBD-B immature oocytes has the smallest sample size of total immature oocytes (8.4%; 27/322). The morphological assessment of the nuclear maturation of all

27 oocytes (prior to fixation) showed an absence of first polar bodies and the germinal vesicle.

134 The chromosomal configuration released from these oocytes post-fixation showed non-dispersed, condensed, metaphasic chromosomes (Figure 2.3-D) (Figure 1.4), in contrast to the typical dispersed, condensed, metaphasic chromosomes post-oocytes lysing. This group of oocytes has been studied previously, and as accounts also for the minority of the in vitro matured oocytes studied (Vlaisavljevic et al., 2007). This group of oocytes have been observed also by other group (Pellestor & Sele, 1988) in which Thirteen out of 188 unfertilized oocytes did not exhibit the first polar body and were found to be in the first-metaphase stage. Vlaisavljevic and his group suggest that the first meiotic division might have occurred resulting in tetraploids - with no chromosomal material extruded with the first polar body, and similar to the present study. About

63% (17/27) out of the total of these oocytes showed uniformly four FISH signals per chromosome analyzed. Whereas, the remaining nine oocytes displayed four FISH signals for the majority of chromosomes analyzed and one chromosome had an extra or missing FISH signal.

As only two or six signals may represent a monosomy or trisomy (similar to immature oocyte at this stage), estimating the incidence of the chromosomally abnormal immature oocytes at this stage can be misleading due to possible confusion with first meiotic errors.

Interpreting FISH signals in immature oocytes at different maturation stages is complex, and estimating the incidence of chromosomally abnormal immature oocytes may overestimate the overall rate of pre-meiotic mitotic errors. Rowsey and his group (2013) have not only excluded foetal oocytes at other stages than leptotene, but have also calculated the incidence of the chromosomally normal oocytes (Rowsey et al., 2013). Therefore, we estimated the incidence of chromosomally normal immature oocytes, as well as chromosomally normal mature oocytes in relation to maternal age.

135

Figure 1.4: The chromosomal configuration and FISH signals pattern of immature oocytes post GVBD-B stage. The image taken by light microscopy (left) shows the chromosomal configuration of immature oocytes post germinal vesicle break down group B (GVBD-B) as intact, metaphasic chromosomes. The image taken by fluorescence microscopy (middle; our study) shows the FISH signals pattern as uniformly tetrad signals per chromosome; four green signals for chromosome 15 and four orange signals for chromosome X. The image taken by fluorescence microscopy (right; (Vlaisavljevic et al., 2007) shows the FISH signals pattern as uniformly tetrad signals per chromosome; four red signals for chromosome 13, four aqua signals for chromosome 16, four blue signals for chromosome 18, four green signals for chromosome 21, and four gold signals for chromosome 22.

136 4.3. Estimated Rates of Chromosomally Normal Immature and Mature Oocytes

The Estimated Rates of Chromosomally Normal Immature and Mature Oocytes in Association to

Maternal Age

Due to the possibility of misdiagnosing the chromosomally abnormal immature oocytes using FISH analysis, the third objective of this project aimed to estimate the overall proportion of chromosomally normal immature oocytes. This would accurately, indirectly determine the association between the incidence of pre-meiotic mitotic errors and maternal age.

The overall estimated rates of chromosomally normal immature and mature oocytes

We found that over half of the immature oocytes of each group were chromosomally normal; the overall proportion of chromosomally normal immature oocytes at the GV and MI stages was estimated to be 73% (72/92) and 53% (28/53) respectively. Sixty three % (17/27) of immature oocytes post the GVBD-B stage were chromosomally normal. These results were consistent with those in a similar study conducted on foetal ovarian oocytes, where the authors determined the proportion of chromosomally normal oocytes at the leptotene stage to be 66.3% for chromosome 16 and 62.3% for chromosome 13 and 21 in all oocytes analyzed (Rowsey et al., 2013). In contrast, 50.7% (34/67) and 37.1% (52/140) of all successfully analyzed mature oocytes at the MII stage and unfertilized oocytes, respectively, were chromosomally normal.

These results are also in agreement with several studies performed on mature oocytes originating from a similar range of maternal ages (Gabriel et al., 2011; Magli et al., 2006; Pujol et al., 2003;

Vlaisavljevic et al., 2007). Taken together, these results strongly underscore the recurrence of first meiotic errors compared to pre-meiotic mitotic errors.

137 Maternal-age related rates of chromosomally normal immature and mature oocytes

In order to determine whether a potential maternal-age related effect on the overall rates of the chromosomal abnormalities in mature and immature oocytes, we first calculated the proportion of chromosomally normal immature oocytes in each maternal age group. Our results indicate that 67.7% (42/62) and 73.3% (22/30) of immature oocytes at the GV stage were normal in younger and older women, respectively. Interestingly, there was no significant difference between the two maternal age groups in the occurrence of chromosomal abnormalities. Similarly,

57.9% (22/38) and 40% (6/15) of immature oocytes post the GVBD-A stage (or at the MI stage, as suggested) were chromosomally normal in both age groups, with no significant difference between them (Table 3.6). These comparable rates suggest that pre-meiotic mitotic errors may not be maternal-age. Using CGH array analysis, similar conclusions were obtained from the study of adult immature oocytes from women who underwent COH treatment cycles (Daina et al., 2014). Thus, our results as well as those from (Daina et al., 2014) are in contrast to the proposed OMSM model by (Hulten et al., 2008) which justified the maternal age effect on the increased rate of pre-meiotic mitotic errors by suggesting that a selection might occur against the trisomic immature oocytes for maturation for further ovulation. This, they proposed would result in an increase in these trisomic oocytes in the ovarian reserve of women with advance maternal age, compared to younger women. Future studies in this regard will immensely benefit from including a larger sample size and coupling DNA analysis techniques with FISH.in adult immature oocytes.

Interestingly, we observed a statistically significant (P < 0.0001) difference in the proportion of chromosomally normal mature oocytes at the MII stage, between the two age groups. In agreement with this, 24 out of the total number of unfertilized oocytes (61.5%)

138 derived from the younger women’s group and only ten from the older women were chromosomally normal, once again being significantly different (P=0.037) (Table 3.6). Such association is well documented in human mature oocytes (Angell, 1994; Dailey et al., 1996;

Fragouli, Wells, Whalley, et al., 2006; Gabriel et al., 2011; Golbus, 1983; Handyside et al., 2012;

T. Hassold & Chiu, 1985; T. J. Hassold & Jacobs, 1984; Kuliev et al., 2011; Nakaoka et al.,

1998; Pellestor et al., 2003; Zenzes et al., 1992) although the underlying molecular basis remains unclear. Given the high clinical relevance of this issue, many hypotheses have been proposed to explain the effect of maternal age on the increased rate of chromosome abnormalities. Among of these hypotheses, two models have been studied extensively: the complete production line model and the prolonged meiotic arrest model. The production line model is the oldest postulated hypothesis, following observations of a significant increase in the frequency of univalents in mature oocytes derived from old mice compared to those derived from young mice (Henderson

& Edwards, 1968). Henderson and colleagues suggested that the increased rate of univalents is maternally age related. These univalents are attributed to bivalents that lack chiasmata (termed achiasmatic), or, alternatively, bivalents with a single chiasma that is either distal or proximal to the centromere in germ cells. They reasoned that since recombination takes place during early oogenesis in the foetal ovaries, the last pool of oogonia to enter meiosis are the last to be ovulated as secondary oocytes, which undergo inadequate recombination. Therefore, the last oocytes to be ovulated are the most susceptible to nondisjunction. Many observations have been reported in order to examine the complete production line model; however, only conflicting results have been obtained (Coop et al., 2008; Hussin et al., 2011; Kong et al., 2004; Polani &

Crolla, 1991; Rowsey et al., 2014).

139 The meiotic arrest model was proposed following discovery of the significant depletion of cohesions in mature oocytes derived from old mice compared to those derived from young mice

(Chiang et al., 2010; Hodges et al., 2005; Lister et al., 2010; L. Liu & Keefe, 2008; Tachibana-

Konwalski et al., 2010). Similar findings have been reported in human oocytes in relation to maternal age (Duncan et al., 2012; Garcia-Cruz, Brieno, et al., 2010; Tsutsumi et al., 2014). It has been found that the cohesion complexes in meiosis have dual functions in binding the homolog around the distal arm regions, and the sister chromatids around the centromere regions for proper segregation at first and second meiotic divisions, respectively. Although these complexes are loaded during DNA synthesis in early foetal development, cohesions around the centromere regions are found to be more degraded in oocytes derived from older women compared to their counterparts derived from younger women. Moreover, the levels of Rec8 and

Smc1 have been found to be decreased significantly in women in their forties compared to those in their twenties. Such depletion contributes to the premature separation of homologs and/or sister chromatids, resulting in first meiotic errors.

In our analysis, we found 81% (13/16) and 67% (14/21) of chromosomally normal immature oocytes post the GVBD-B stage that showed four FISH signals for all chromosomes analyzed, with no significant difference between both maternal age groups (Table 3.6). It has been suggested that the first meiotic division took place in this group of oocytes where a non- disjunction occurred for all chromosomes. This resulted in tetrads confined to only the mature oocytes, an observation that has been previously documented (Vlaisavljevic et al., 2007).

Accordingly, these tetrads should be classified as first meiotic errors, but they indirectly indicate chromosomally normal precursor immature oocytes where four signals were shown per chromosome analyzed, while two and four signals represent a monosomy or trisomy of the

140 precursor immature oocytes, respectively. Although there was no significant difference between the maternal age groups, the sample size of this group is not large enough to statistically analyze for an association. Therefore, the FISH signals from this group should be interpreted with caution, until a larger sample size is achieved.

The Estimated Rates of Chromosomally Normal Immature and Mature Oocytes in the Siblings

Study Group in Association to Maternal Age

In our analysis of sibling oocytes, we re-calculated the overall rate of chromosomally normal oocyte immature and mature oocytes where each patient donated a minimum of one immature oocyte and one mature oocyte. In order to eliminate the possible effect of patient-to- patient variation on the overall occurrence of chromosomal abnormalities, only cohorts of oocytes received on the day of analysis (i.e. immature oocytes at any maturation stage as well as those matured in vitro) were included in the re-analysis. A total of 49 immature oocytes at any maturation stage and 27 mature oocytes at the MII stage were donated by the younger women’s group A comparable sample size of 37 immature oocytes at any maturation stage but about double the sample size of 50 mature oocytes were derived from the older women (Table 3.7).

Such difference in sample size of donated sibling immature and mature oocytes may account for the statistical significance between the two age groups (p=0.003) (Table 3.7) (Figure 3.4).

Effect of maternal age on rates of chromosomal abnormalities in immature and mature oocytes

The present study reports for the first time, the estimated proportion of chromosomally normal immature and mature oocytes among sibling oocytes. All results obtained from this cohort of oocytes are in agreement with results obtained from the analysis of the non-sibling oocytes. This suggests that patient-to-patient variation may not have a major effect on the overall estimated rates of chromosomal abnormalities in oocytes. .

141 4.4. Estimated Rates of Pre-Meiotic Mitotic Errors in Mature Oocytes

Older studies considered the presence of an extra, non-complementary FISH signal in the analyzed oocyte as an artefact (Dailey et al., 1996; Munne et al., 1995). Further observations on the incidence of pre-meiotic mitotic errors in mature oocytes and their corresponding first polar bodies have been reported (Cupisti et al., 2003; Fragouli, Wells, Whalley, et al., 2006; Garcia-

Cruz, Casanovas, et al., 2010; Gutierrez-Mateo, Benet, et al., 2004; Pujol et al., 2003). However, these studieswere limited in their scope to analyze this type of chromosomal abnormality, as they primarily targeted the incidence of first meiotic errors and its relation to maternal age using FISH and CGH techniques. Only two studies (Mahmood et al., 2000; Obradors et al., 2010) and a case report (Cozzi et al., 1999) were designed to investigate the incidence of pre-meiotic mitotic errors in doublets of mature oocytes and their corresponding first polar bodies.

The overall rate of pre-meiotic mitotic errors in mature oocytes

Our study reports a low rate of chromosomal abnormalities in mature oocytes due to pre- meiotic mitotic errors, which is in agreement with previously reported overall rates in mature

(Obradors et al., 2010) and immature oocytes (Daina et al., 2014). In trisomic precursor immature oocytes, we noted that they are 50% likely to result in a chromosomally abnormal mature oocyte due to pre-meiotic mitotic errors in one scenario, and 100% likely to give rise to a chromosomally abnormal mature oocyte due to pre-meiotic mitotic errors in the other scenario.

In mature oocytes on the other hand, our results are inconsistent with the previously documented observations on the incidence of pre-meiotic mitotic errors (Cupisti et al., 2003;

Fragouli, Wells, Whalley, et al., 2006; Garcia-Cruz, Casanovas, et al., 2010; Gutierrez-Mateo,

Benet, et al., 2004; Pujol et al., 2003). Nevertheless, it is important to note that these studies were not intended to analyze this particular type of chromosome abnormality. In fact, these studies

142 were designed to develop and standardize a larger panel of FISH probes and CGH techniques for chromosomal screening in mature oocytes and or their corresponding first polar bodies.

Therefore, a higher rate of experimental artefacts was expected. Also, a relatively small sample size of mature oocytes with the corresponding first polar bodies and mixed population of oocytes

(unfertilized and in vitro matured oocytes whether subjected to sperms) were analyzed. This may explain the high discrepancy in the estimated incidence of pre-meiotic mitotic errors in these cells. In sum, the present study reports the overall rate of pre-meiotic mitotic errors in the largest sample size of mature oocytes studied to date, allowing us to reasonably conclude that the contribution of pre-meiotic mitotic errors to the overall rate of chromosome abnormalities in human is considerably low.

Patterns of chromosome segregation involved in pre-meiotic mitotic errors

We were able to observe both proposed patterns of the extra chromosome due to pre- meiotic mitotic errors at anaphase I stage as represented in (Figure 1.6, 3.5). In the first pattern, a double gain is observed of the screened chromosome in one of the doublets cells (MII-1PB)

(Four FISH signals in either cell) and the other cell was diploid of the same chromosome. In the second pattern, a gain of the screened chromosomes is observed in both doublets cells as a chromatid per cell (MII-1PB) (Three FISH signals per cell). We found a two-fold increase in the occurrence of pre-meiotic mitotic errors displaying the first pattern in comparison to those exhibiting the second pattern of pre-meiotic mitotic errors (n=21 vs. n=7 (14), respectively)

(Table 3.9). This pattern has been reported frequently among the limited studies (Cozzi et al.,

1999; Cupisti et al., 2003; Mahmood et al., 2000; Obradors et al., 2010; Pujol et al., 2003)

(Figure 2.4).

143

Figure 2.4: Non-disjunction of the extra chromosome due to pre-meiotic mitotic error in either cell of MII-1PB doublet. Two extra, non-complementary FISH signals in mature oocyte (Left; our study) or in first polar body (right; (Mahmood et al., 2000), while the corresponding cell is diploid of the same chromosome.

144 Other studies conducted on foetal oocytes have demonstrated that the incidence of forming a trivalent in trisomic oocytes at the prophase I stage is more greater than that of forming a bivalent and univalent in trisomic oocytes for chromosomes 13, 16, 21, and 18 (Barlow et al.,

2002; E. Y. Cheng et al., 2009; Hartshorne, Barlow, Child, Barlow, & Hulten, 1999; Hulten et al., 2008; Robles et al., 2007; Roig, Robles, Garcia, Martinez-Flores, et al., 2005; Speed, 1984,

1985; Tease et al., 2006; Tease et al., 2002). One study proposed that the paired chromosome as a trivalent may segregate in a non-disjunction fashion, resulting in a double gain of the chromosomes in one of the doublets cells (MII-1PB) while the other cell remains diploid for the same chromosome at the metaphase I stage. The paired chromosome as a bivalent with a univalent may segregate in a PD fashion, resulting in a gain of a chromatid of the chromosome involved in both doublets cells (MII-1PB) (PD leading mechanism) (figure 1.6, 1.1) (Hulten et al., 2008). Such patterns of the extra chromosome segregation have been observed in other studies that analyzed the chromosomal complements in adult mature oocytes (Cupisti et al.,

2003; Mahmood et al., 2000; Obradors et al., 2010; Pujol et al., 2003) using FISH and CGH- based analyses These studies suggested that the trivalent pattern of homologue pairing tends to progress more quickly in shorter chromosomes than in longer ones during human female prophase, whereas the trivalents that are formed at the pachytene stage are well maintained until the diplotene stage (Robles et al., 2007; Roig, Robles, Garcia, Martin, et al., 2005; Roig, Robles,

Garcia, Martinez-Flores, et al., 2005).

Chromosomes involved in pre-meiotic mitotic errors

With the exception of the X chromosome, our study found that all the autosomes analyzed were involved in pre-meiotic mitotic errors in mature oocytes and their corresponding polar bodies. The highest rate of errors was found for chromosome 13, followed by 16, 18 and 22 as

145 shown in Table (3.9). Our results re in agreement with both studies conducted on mature and immature oocytes (Daina et al., 2014; Obradors et al., 2010; Pujol et al., 2003). This suggests that these extra, smaller in size chromosomes in trivalent may not only well maintained till the diplotene stage, but also may able to reach the metaphase II stge.

Overall rate of pre-meiotic mitotic errors in association with maternal age

We found that the estimated rates of pre-meiotic mitotic errors were comparable in mature oocytes, including unfertilized oocytes derived from younger and older women groups (9.1%;

9/99 vs. 13.4%; 15/108, respectively) with no significant differences between the maternal age groups (Table 3.9). These results as well as those that were estimated in immature oocytes are in agreement with previously reported findings in mature and immature oocytes (Daina et al., 2014;

Obradors et al., 2010; Pujol et al., 2003), suggesting that pre-meiotic mitotic errors are not maternally age dependent as first meiotic errors may be.

Overall rate of pre-meiotic mitotic errors in mature oocytes

As there was no association between the incidence of pre-meiotic mitotic errors and advanced maternal age, we attempted to determine whether there is an association between the incidence of these types of errors and female infertility. We found 17 out of 112 (13.9%) patients who underwent COH treatment cycles carried pre-meiotic mitotic errors. Among of these patients, three donated more than one oocyte with pre-meiotic mitotic errors (Table 3.9). All of these patients had a history with primary infertility, except for four who attended the fertility clinic due to secondary infertility. Nine and two out of 17 patients had a history with RPL and 1-

2 miscarriages (MC) donated oocytes with pre-meiotic errors, respectively, where two out of

RPL patients were translocation carriers as indicated in Table 3.9. As most of these patients donated only oocytes with pre-meiotic errors, it was difficult to detect a true association between

146 the incidence of these types of errors and miscarriage using FISH. In fact, the association between patients with recurrent miscarriage and gonadal mosaicism has been well demonstrated in ovarian cells and mature oocytes among translocation carriers as well as in younger women with multiple trisomic conceptions (Bruyere et al., 2000; Cozzi et al., 1999; Hulten et al., 2008;

Sachs et al., 1990).

It has been suggested to reserve the terminology of gonadal mosaicism to errors when the same chromosome is involved in a proportion of germ cells or gametes, resulting in multiple conceptions. These errors have been postulated to arise as mitotic errors occurring during preimplantation embryonic development to further get confined to germline of the same embryo during post implantation embryonic development. However, the terminology of germline mosaicism should be reserved for errors when different chromosomes are involved in a proportion of germ cells or gametes which are more likely to arise during the proliferative stage

(Delhanty, 2011).

Estimated Rates of First Meiotic Errors in Mature Oocytes

Overall rate of first meiotic errors in mature oocyte, including unfertilized oocytes

We found a considerably high incidence of first meiotic errors (92.5%) in mature oocytes and (58.5%) in unfertilized oocytes with the corresponding first polar bodies, with high correlation (r = 0.81). These chromosomally abnormal oocytes with errors were associated with maternal age (Figure 3.7) confirming all previously reported data using different methodologies for chromosomal analysis (Angell, 1994; Dailey et al., 1996; Fragouli, Wells, Whalley, et al.,

2006; Gabriel et al., 2011; Golbus, 1983; Handyside et al., 2012; T. Hassold & Chiu, 1985;

Kuliev et al., 2011; Nakaoka et al., 1998; Pellestor et al., 2003; Zenzes et al., 1992). Moreover, our results confirmed the findings of these studies where premature division (PD) is not only the

147 main mechanism leading to complementary first meiotic errors in both maternal age groups compared to AL and ND, but also the most predominant mechanism leading to the higher incidence of chromatid errors in oocytes from older women compared to those from younger group compared to AL. These results are in agreement with the possibility of more depleted cohesions in oocytes retrieved from older women compared from younger group (Duncan et al.,

2012; Garcia-Cruz, Brieno, et al., 2010; Tsutsumi et al., 2014).

In addition, we found that first meiotic errors for chromosome 13 and 21 were the highest in oocytes derived from both maternal age groups, followed by chromosome 15, 16 and 22 and chromosome 18 and X (Gabriel et al., 2011; Handyside et al., 2012; Kuliev et al., 2011).

Overall rate of first meiotic errors in association with in vitro maturation

Although the present study was not designed to determine whether there is an association between the incidence of first meiotic errors and in vitro culture, we analyzed the chromosomal complements of in vitro matured oocytes and unfertilized oocytes collected from only younger women as separate groups. The rationale behind this analysis was to determine whether the incidence of first meiotic errors was affected in in vitro cultured oocytes. Our results showed that there was no significant difference between the two groups. Similar findings were obtained when we calculated the difference between the in vitro matured oocytes collected at Day 1 and Day 3 following egg retrieval. This may be explained by the fact that these oocytes might be matured around the same time but received later for analysis (Pujol et al., 2003; Vlaisavljevic et al.,

2007).

4.5. Limitations

The major limitation in our study is using FISH analysis as a stand-alone methodology for analysis of chromosomal complement in oocytes. Immunofluorescence (IF) analysis may, for

148 instance, confirm our resulted obtained from immature oocytes at all maturation stages , given the considerable variation in FISH signals for the chromosomes analyzed in this group Studies have successfully demonstrated the the incidence of pre-meiotic mitotic errors in foetal ovarian cells using a combination of FISH and IF techniques (Hulten et al., 2008; Morris et al., 2012;

Robles et al., 2007; Roig, Robles, Garcia, Martin, et al., 2005; Rowsey et al., 2013). In these studies, the exceptionally large number of ovarian cells allowed them to develop and validate both techniques since cell loss was not an issue. However, in our case, donated samples were very difficult to obtain and we attempted to overcome this issue by validating FISH signals statistically, and calculating the chromosomally normal oocytes to estimate indirectly the incidence of pre-meiotic mitotic errors and its association to maternal age.

Another limitation of our study is the small sample size. Because the incidence of pre- meiotic mitotic errors is considerably low compared to first meiotic errors, increasing the sample size is required to improve the statistical power of the test. Although our study represented the largest sample size of successfully analyzed immature and mature oocytes with the corresponding first polar bodies compared to other groups (Cupisti et al., 2003; Daina et al.,

2014; Obradors et al., 2010; Pujol et al., 2003), more donated samples per patients are needed in order to confirm the clinical relevance of pre-meiotic mitotic errors and first meiotic errors in oocytes.

149

CHAPTER FIVE

CONCLUSION AND FUTURE DIRECTION

150 5.1. Conclusion

In the present study, we have analyzed the chromosomal complements of human immature and mature oocytes derived from women who underwent COH treatment cycles in association to maternal age using FISH-based analysis for those chromosomes that are most commonly involved in pregnancy loss. To our knowledge, this study is the first to analyze the chromosomal complements using such a large sample size of human immature oocytes (at different maturation stages), and mature oocytes with the corresponding first polar bodies using FISH analysis.

We demonstrated that:

1. The overall rates of chromosomally normal immature oocytes were comparable at all

maturation stages: immature oocytes at GV stage, immature oocytes post GVBD-A stage

(MI stage as suggested), and immature oocytes post GVBD-B stage with no association to

maternal age compared to chromosomally normal mature oocytes.

2. The reanalyzed overall rates of chromosomally normal immature and mature oocytes of the

same cohort (sibling’s oocytes) confirmed the similar overall rates of chromosomally

normal immature oocytes with no association to maternal age compared to chromosomally

normal mature oocytes.

3. The incidence of pre-meiotic mitotic errors was considerably low in mature oocytes with

the corresponding first polar bodies compared to first meiotic errors. There was no

association between maternal age and the overall rate of pre-meiotic mitotic errors in

mature oocytes compared to first meiotic errors.

All together, we concluded that incidence of pre-meiotic mitotic errors in human immature and mature oocytes derived from women who underwent COH treatment cycles is relatively low compared to first meiotic errors. This suggests that the contribution of these errors to the overall

151 rate of maternally related chromosome abnormalities is considerably low, but not trivial. Also, there is no association between advanced maternal age and pre-meiotic mitotic errors compared to first meiotic errors. This suggests that these errors are not maternal-age dependent as first meiotic errors; thus, our findings reject the proposed Oocyte Mosaicism Selection Model

(OMSM) by (Hulten et al., 2008). Last, pre-meiotic mitotic errors may not have a significant negative effect on human reproduction; however, consequences of these errors have clinical relevance to human fertility, at least in our study population.

5.2. The Study Future Direction

As previously mentioned, pre-meiotic mitotic errors arise as mitotic errors during the proliferative stage in very early oogenesis, resulting in chromosomally abnormal foetal oocyte.

Any of the foetal oocytes may ultimately become immature oocyte in the adulthood. Pre-meiotic mitotic errors can also arise as mitotic errors during the preimplantation embryo development, resulting in chromosomally abnormal cells. These cells may ultimately become precursor to germ cells that carry the same chromosomal error (Delhanty, 2011).

Hypothetically, different chromosomes might be involved in the chromosomally abnormal germ cells due to pre-meiotic mitotic errors, as have been previously suggested (Cupisti et al.,

2003; Daina et al., 2014; Mahmood et al., 2000; Morris et al., 2012; Obradors et al., 2010; Pujol et al., 2003; Rowsey et al., 2013). On the other hand, the same chromosome can be involved in the resulting abnormal germ cells which are generated from mitotic errors occurred during preimplantation embryonic development of the same individual, as has been previously reported

(Cozzi et al., 1999). In this case report, the author investigated the origin of trisomy 21 in unfertilized oocytes and preimplantation embryos derived from young woman had a history with multiple conceptions with trisomy 21 who underwent IVF/PGS treatment cycle. The cytogenetic

152 analysis showed a high incidence of aneuploid unfertilized oocytes and preimplantation embryos with trisomy 21 (five out of seven preimplantation embryos and three out of four unfertilized oocytes). Also, the molecular analysis revealed that the extra chromosome 21 is maternally inherited, suggesting that trisomic germ cells are present within the patient’s gem cell line.

Accordingly, we propose to investigate the origin of pre-meiotic mitotic errors in human mature and or mature oocytes and the incidence of these errors occurred by either scenario. This would not only determine the overall rate of pre-meiotic mitotic in human oocytes, but also identify for first time the incidence of these errors in relation to their origin.

As it could be difficult to obtain several immature and or mature oocytes per patient for research, and only chromosomal complements of these oocytes were analyzed using FISH technique, we propose the following:

Array-based analysis for the chromosomal complements in donated human immature and mature oocytes

aCGH-based analysis is well developed for a comprehensive chromosomal screening in human mature oocytes compared to SNP-based array analysis (Capalbo et al., 2013;

Christopikou et al., 2013; Fragouli et al., 2013; Gabriel et al., 2011; Geraedts et al., 2011;

Handyside et al., 2012), we suggest applying this technique to comprehensively investigate the incidence of pre-meiotic mitotic errors in human oocytes.

PCR-based analysis for DNA polymorphism in parental lymphocytes or buccal cells

The whole genome amplified DNA of chromosomally abnormal oocyte due to pre-meiotic mitotic errors, as well as the whole genome amplified DNA of lymphocytes (parental and patients peripheral blood samples) can be analyzed by PCR-based technique using STR markers to identify the origin of pre-meiotic mitotic errors, as previously described (Cozzi et al., 1999).

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