Involvement of Nlrp5 in the Maintenance of Genome Integrity in Murine Oocytes

INVOLVEMENT OF NLRP5 IN THE MAINTENANCE OF GENOME INTEGRITY IN MURINE OOCYTES

RUSSANTHY VELUMMAILUM

A thesis submitted in conformity with the requirements for the degree of Master of Science Graduate Department of Physiology University of Toronto

INVOLVEMENT OF NLRP5 IN THE MAINTENANCE OF

GENOME INTEGRITY IN MURINE OOCYTES

RUSSANTHY VELUMMAILUM

Master of Science

Graduate Department of Physiology

University of Toronto

2011 ABSTRACT

Nlrp5, a maternal-effect gene, is required for embryonic progression and female fertility in mice. Previous work indicated an age-related decline in Nlrp5 transcripts in murine oocytes. As maternal age is associated with increased spindle organization defects, studies in this thesis focused on the analysis of meiotic spindle defects in oocytes of Nlrp5-deficient mice. NALP5 protein showed a novel kinetochore-localization pattern, which was disturbed by spindle poisons. Nlrp5-deficient oocytes displayed a higher frequency of spindle abnormalities and chromosomal misalignment. Upon fertilization, these defects translated into increased incidences of multinucleation. As these phenotypes are associated with deficiencies in genome stability, we examined spindle assembly checkpoint (SAC) components. We found that numerous SAC proteins were dysregulated, implying that NALP5 may be critical in sensing oocyte-related SAC defects. We found that Nlrp5-deficient oocytes may have increased DNA damage. Thus, Nlrp5 may be an integral component responsible for preservation of genome integrity in female gametes.

ACKNOWLEDGEMENTS

I would like to express heartfelt gratitude to my supervisor, Dr. Andrea Jurisicova, for providing me with the scientific insight and guidance for my Master’s thesis. I am especially thankful for this graduate studies opportunity.

I wish to acknowledge my supervisory committee members, Dr. Anthony Gramolini and Dr. Susannah Varmuza, for all of their advice and their willingness to guide my project.

I specially want to thank Dr. Alagammal Perumalsamy for her real time PCR work on p73.

I also want to thank all members of the Jurisicova lab, including: Dr. Jacqui Detmar, Dr. Alagammal Perumalsamy, Dr. Han Li, Aluet Borrego-Alvarez, Shakib Omari, Tania Yavorska, Taline Naranian, Jasmine Chong, Roxanne Fernandes, and Fatima Khan. I also want to extend a heartfelt thanks to all members of the Samuel Lunenfeld Research Institute’s sixth floor for your friendship. All of you have helped make my Master’s experience a treasurable one!

Finally, I would like to express my sincerest gratitude to my family and friends for all of their support. I would like to dedicate this thesis to my mother, Shantha Ruthiran, and my father, Ruthiran Velummailum, for their unconditional love and continued encouragement throughout my Master’s career. Thank you!

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TABLE OF CONTENTS

Abstract ...... ii Acknowledgements...... iii Table of Contents ...... iv List of Tables ...... vii List of Figures ...... viii List of Abbreviations ...... ix 1 Introduction ...... 1 1.1 Infertility ...... 1 1.2 Aneuploidy ...... 3 1.3 Oogenesis ...... 4 1.4 Early Mammalian Embryonic Development ...... 6 1.5 Maternal-Effect Factors ...... 9 1.6 NALP5/MATER ...... 11 1.6.1 Structural Domains and NALP Family Members ...... 11 1.6.2 Identification of NALP5 ...... 12 1.6.3 NALP5 Expression Pattern ...... 13 1.6.4 Nlrp5-deficient Phenotype ...... 14 1.6.5 NALP5 Interacting Partners ...... 15 1.7 Spindle Assembly Checkpoint ...... 17 1.8 DNA damage ...... 26 1.9 Rationale ...... 30 Hypotheses ...... 31 Objectives ...... 31 2 Methods ...... 32 2.1 Mouse Husbandry ...... 32 2.2 Nlrp5 Genotyping ...... 32 2.3 Oocyte/Zygote Retrieval ...... 33 2.3.1 Metaphase II (MII) Oocyte Collection ...... 33

2.3.2 Germinal Vesicle (GV) Oocyte Collection...... 33 2.3.3 Zygote (Fertilized oocyte) Collection ...... 34 2.4 Whole-Mount Oocyte Immunocytochemistry ...... 34 2.4.1 Negative Controls and Blocking Peptide ...... 37 2.5 Nocodazole Treatment ...... 37 2.6 UV irradiation and Oocytes Culture ...... 38 2.7 Comet Assay ...... 38 2.8 Imaging: Fluorescence Microscopy ...... 39 2.9 Microarray ...... 40 2.10 Statistical Analysis...... 40 3 Results ...... 42 3.1 Histone Methylation Pattern in Nlrp5-Deficient Zygotes ...... 42 3.2 The role of Nlrp5 in the activation of the Spindle Assembly Checkpoint in Metaphase II Oocytes ...... 44 3.2.1 Pronuclear Number Abnormalities in Nlrp5-deficient Zygotes ...... 44 3.2.2 Spindle Morphology and Chromosomal Misalignment Abnormalities in Ovulated, Metaphase II Oocytes...... 46 3.2.3 NALP5 Localization in MII Oocytes ...... 50 3.2.4 Spindle Assembly Checkpoint Analysis in Nlrp5 Deficiency at the MII Oocyte-stage ...... 52 3.2.4.1 BUB1 Protein Expression ...... 52 3.2.4.2 BUBR1 Protein Expression ...... 54 3.2.4.3 p73 and TAp73 Protein Expression ...... 56 3.2.4.4 Aurora kinase B ...... 58 3.2.4.5 Aurora kinase C ...... 58 3.2.4.6 Dynactin p50 Protein Expression ...... 59 3.2.5 Nocodazole Treatment in Metaphase II Oocytes ...... 62 3.3 The role of Nlrp5 in triggering an appropriate DNA damage response in oocytes ...... 64 3.3.1 UV Irradiation Response in MII Oocytes ...... 64 3.3.1.1 NALP5 and UV Damage in MII Oocytes...... 65 3.3.1.2 Phosphorylated Histone H2AX (γH2AX) ...... 67

3.3.1.3 53BP1 ...... 69 3.3.1.4 Phosphorylated Histone H3 at Serine 10 (H3S10) ...... 71 3.3.1.5 BRCA1 ...... 73 3.3.1.6 ATM ...... 75 3.3.2 Comet Assay in MII Oocytes with Nlrp5 Deficiency ...... 75 3.3.3 UV Damage and α-Tubulin ...... 77 3.3.4 DNA damage markers and Nocodazole ...... 78 3.4 Microarray Results...... 80 4 Limitations ...... 81 5 Discussion ...... 83 5.1 Spindle Assembly Checkpoint ...... 86 5.2 DNA Damage ...... 89 6 Future Directions ...... 93 7 References ...... 95 8 Appendix ...... 106

LIST OF TABLES

Table 1: Examples of Maternal-Effect Genes and their Phenotypes ...... 10 Table 2: List of Spindle Assembly Checkpoint Proteins and their Phenotypes ...... 23 Table 3: Frequency of Spindle Configuration and Chromosomal Segregation Defects ...... 25 Table 4: List of Primary Antibodies and Dilutions ...... 36 Table 5: List of Secondary Antibodies and Dilutions ...... 37 Table 6: mRNA transcripts with Decreased Expression in Nlrp5-Deficient MII oocytes ...... 80 Table 7: mRNA transcripts with Increased Expression in Nlrp5-Deficient MII oocytes ...... 80 Table 8: List of Antibodies that did not work...... 82 Table 9: Complete list of mRNA transcripts with Decreased Expression in Nlrp5-Deficient MII oocytes ...... 106 Table 10: Complete list of mRNA transcripts with Increased Expression in Nlrp5-Deficient MII oocytes ...... 111

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LIST OF FIGURES

Figure 1: Oogenesis...... 6 Figure 2: Early Mammalian Embryonic Development...... 8 Figure 3: Mouse and Human NALP5 protein structure...... 12 Figure 4: Subcortical Maternal Complex...... 16 Figure 5: Metaphase II-arrested meiotic spindles ...... 17 Figure 6: Spindle Assembly Checkpoint mode of action...... 18 Figure 7: Aurora kinase pathway...... 21 Figure 8: DNA Damage/Repair Response Pathway in Somatic cells...... 27 Figure 9: Methylation pattern in Early and Late Nlrp5-deficient zygotes...... 43 Figure 10: Pronuclear Abnormalities in Nlrp5-deficient zygotes...... 45 Figure 11: Proportion of ovulated oocytes with abnormal spindle morphology...... 47 Figure 12: Proportion of ovulated oocytes with misaligned chromosomes...... 49 Figure 13: Localization of NALP5 protein in MII oocytes...... 51 Figure 14: BUB1 Protein expression in MII oocytes...... 53 Figure 15: BUBR1 Protein expression in MII oocytes...... 55 Figure 16: p73 and TAp73 expression pattern in MII oocytes...... 57 Figure 17: Relative expression of various Spindle Assembly Checkpoint proteins in wildtype and Nlrp5-deficient MII oocytes...... 60 Figure 18: NALP5 expression with Nocodazole treatment in MII oocytes...... 63 Figure 19: NALP5 expression after UV exposure...... 66 Figure 20: γH2AX expression after UV exposure and in Nlrp5 deficiency...... 68 Figure 21: 53BP1 expression after UV exposure and in Nlrp5 deficiency...... 70 Figure 22: H3S10 expression after UV exposure and in Nlrp5 deficiency...... 72 Figure 23: BRCA1 expression after UV exposure and in Nlrp5 deficiency...... 74 Figure 24: Comet assay in MII oocytes...... 76 Figure 25: α-Tubulin localization after UV exposure...... 77 Figure 26: 53BP1 and γH2AX expression after Nocodazole treatment...... 79 Figure 27: UV damage pathway in wildtype metaphase II-arrested oocytes...... 90 Figure 28: Summary of the spindle and DNA damage-implicated roles of Nlrp5 in MII oocytes.92

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LIST OF ABBREVIATIONS

DMSO – dimethyl sulfoxide

GV – Germinal Vesicle

H3K9me3 – tri-methylation of histone H3 at lysine 9 hCG – human Chorionic Gonadotropin hpf – hours post fertilization

ICCH – Immunocytochemistry

KO – knockout; Nlrp5-deficient

MI – metaphase of meiosis I

MII oocyte – metaphase of meiosis II-arrested oocyte; ovulated oocyte; mature oocyte

NALP5 – mouse protein

Nlrp5 – mouse Nalp5 gene

PB – polar body

PIPES – 1, 4-Piperazinediethanesulfonic acid

RFU – relative fluorescence units

RT-PCR – Reverse transcription polymerase chain reaction

SAC – Spindle Assembly Checkpoint

UV – Ultraviolet

WT – wildtype; Nlrp5-present

ix 1

1 INTRODUCTION

1.1 INFERTILITY In Canada, an estimated 8.5% of couples of reproductive age experience infertility, defined as the lack of conception after one year of attempting to get pregnant [1]. Of those couples that experience infertility, in 40% of the cases infertility is related to female reproductive factors, in 30% it is due to male reproductive factors, and in the remaining 30% of cases, either male and female factors or other unknown reasons are responsible [1]. The breakthrough treatment for infertility is assisted reproductive technology (ART), pioneered by the recently acclaimed Nobel laureate Robert Edwards’ work in the in vitro fertilization (IVF) treatment. The first live birth resulting from IVF treatment took place in 1978 and since then the pregnancy success rate has been dramatically improving [2]. Interestingly, approximately 1-3% of all births in industrial countries could be attributed to ART. In mammals, the age-associated decline in fertility is likely caused by females producing eggs also known as oocytes that have decreased developmental competence [3]. Reproductive aging is also a major contributing factor to ART failure. In 2007, 37% of ART cycles performed in Canada on women less than 35 years of age resulted in successful pregnancies, while women aged 35-39 had a pregnancy success rate of 27% and women over 40 years old experienced a rate of only 11% [4]. Furthermore, additional data from 2007 comparing the percentages of transfers resulting in live births for ART cycles using donor oocytes or using a woman’s own oocytes, among women of different ages found that the likelihood of implantation of the fertilized oocyte was age-dependent. The live birth rate percentage of transfers from women’s own eggs declined with age, while transfers with donor eggs (from young patients aged 20-30), even in women older than 35 years (up to their late forties), remained consistent above 50% [5]. Hence, given that older women who receive oocytes from young donors have similar pregnancy rates as young women, this implies that the rest of the reproductive tract/uterus is functioning well and it is the oocyte quality that is the culprit in decreased fecundity with aging [5]. Many reasons could contribute to the ability to conceive in females. To a large extent, female fertility is determined by the quality of the oocyte as reflected by its ability to undergo meiosis, to be fertilized, and to produce a healthy embryo [6]. Hence, some frequent defects in female fertility include poor fertilization, poor embryo development and implantation, or even

2 inability to carry the pregnancy to term. We are most interested in understanding the molecular determinants of oocyte quality, as the oocyte is the progenitor of preimplantation embryos and the foetus. There is a subset of infertile women who have failed pregnancies after multiple assisted reproductive cycles and this is often attributed to suboptimal embryo quality [7]. Poor quality embryos are characterized by compromised developmental potential and embryo fragmentation, which are thought to originate from poor oocyte quality and, potentially, lack of maternal-effect factors (discussed in section 1.5). As the oocyte contributes the majority of cytoplasm along with half of the nuclear DNA to form the zygote, oocyte quality is an unquestionably significant determinant of embryo quality [8]. It is probable that the patients with repeated assisted reproductive technology failure may have defective maternal factors in the fertilized oocyte resulting in the inability to support early development. Supplementation of normal cytoplasmic protein into the oocytes of these infertile women might restore normal early development and successful pregnancy [9]. This treatment option for low quality embryo development is known as ooplasmic transfer. This procedure involves the insertion of donor oocyte cytoplasm (ooplasm) into patient oocytes [9]. Although ooplasmic transfers appear successful, this procedure could cause genetic alterations, since the offspring will carry mitochondria from both the donor and recipient [9]. Furthermore, in Canada, treatments that alter the foetal genome, including ooplasmic transfer, are prohibited. As a result, in order to create other treatment options, it is necessary to investigate the molecular causes underlying suboptimal oocyte quality. It is becoming increasingly apparent that oocyte quality is a primary factor in the success of assisted reproduction treatments; however very little is known about the molecular characteristics of a high quality oocyte. It is known that in some cases, there is a morphological difference between low and high quality oocytes [10]. Since poor quality oocytes give rise to compromised embryos, investigating the genome integrity of the oocyte may elucidate a molecular understanding behind oocyte quality.

1.2 ANEUPLOIDY A major contributor to infertility is aneuploidy. Aneuploidy is a type of chromosomal deformity in which there is an abnormal number of chromosomes in daughter cells after cell division. In humans, chromosomal anomalies occur at an incidence of about 0.6% in newborns, 6% in still births, and 60% is spontaneous abortions [11]. More specifically, one of the most well documented causes of pregnancy loss is aneuploidy, occurring in about 35% of spontaneous abortions [12]. It is estimated that 20% of human oocytes are aneuploid [12]. Furthermore, meiotic recombination has a significant effect on the genesis of aneuploidy in both females and males [11]. The aetiology of this disorder has been well-attributed to maternal age [11]. Although most chromosome abnormalities are lethal and are lost early in embryonic development, some of the affected foetuses do survive to term, for example those with trisomies 13, 18, 21 and aneuploidy of the sex chromosomes [11]. Individuals who possess these chromosomal anomalies can have mental and physical disabilities, infertility and behavioural problems.

Human egg donation studies indicate it is the oocyte, not the reproductive tract, that is the major cause of infertility, and is responsible for a marked increase in aneuploidies in women 35 years or older [3, 13]. In addition, older women (greater than 40) have a 45% higher likelihood of generating a foetus with chromosomal abnormalities, compared to women younger than 40 [14]. Although the exact molecular basis behind female reproductive defects and aging is unclear, many studies have revealed age-related changes in oocytes. Some of these changes with increased aging include increased rate of chromosomal abnormalities due to spindle organization defects [15], oocyte aneuploidy [15, 16] and changes in gene expression of spindle assembly-related genes [17].

Many studies on human embryos have associated multinucleation with the presence of aneuploidies and mosaicisms [18]. Multinucleated blastomeres (MNB) have been studied in human embryos. Blastomeres are cells of preimplantation embryos. In human studies, researchers found that 44% of embryos from patients undergoing embryo transfers exhibited MNB. MNB are typically attributed to abnormal embryo development, since about 57% of MNBs arrest at the 2-15 cell stage [19]. Furthermore, fluorescence in situ hybridization studies found the presence of multinucleated cells in non-arrested day-2 or -3 human embryos in 74% of cases of extensive mosaicism/polyploidy [20]. Human MNB may occur at anytime during preimplantation development, but are more often present at the 2-cell embryo stage [18].

Another study found that when more than 50% of transferred embryos contained multinucleated blastomeres there was a significant decrease in implantation, clinical pregnancy and live birth rates [21], suggesting that embryos with MNB have poor developmental potential. In addition, human embryos with unevenly sized blastomeres have lower implantation and pregnancy rates and have a higher incidence of aneuploidy and multinucleation [22]. Since many of these aneuploid phenotypes are associated with decreased chromosomal fidelity and could arise from the oocyte, it is useful to study genome stability in oocytes.

1.3 OOGENESIS Oogenesis is the process of development of the female germ cell, otherwise known as the oocyte. At birth, each female mouse has a finite pool of primordial germ cells, which is in prophase of meiosis I and will remain arrested in this stage for the rest of oogenesis unless selected for ovulation [23]. Just days after birth, somatic cells, called granulosa cells, initiate contact with the germ cell and form the primordial follicle [23]. During the prolonged prophase arrest, which can last for years, oocytes are transcriptionally competent, as dictyate chromosomes are accessible to transcriptional machinery [24]. Each oocyte has numerous cytoplasmic projections from granulosa cells that form specialized junctions [25], thus allowing for intricate intercellular communication between the germ cell and the somatic cells. This communication takes place through the presence of gap junctions and allows for the transfer of metabolites and regulatory substances between the two cell types to allow for controlled growth and differentiation of the oocyte [26]. Gap junctions remain in place as the oocytes grow, even after the deposition of the zona pellucida, a glycoprotein layer separating the oocyte from the granulosa cells [25]. Growth from primordial to primary follicle requires the accumulation of maternal products. As the oocyte increases in size, it acquires the competence to enter the final stages of meiosis in response to either the correct hormonal stimulation (in vivo) or release from the follicle (in vitro) [23]. Oocytes will become atretic unless correct levels of pituitary hormones, follicle stimulating hormone (FSH) and luteinizing hormone (LH) are present [25]. FSH causes the stimulation of a few follicles in each cycle, the granulosa cell proliferation and, in turn, the follicle accumulates fluid, swells and pushes toward the periphery of the ovary. At this stage, the follicle is referred to as the Graafian follicle. The expanded Graafian follicle, with a fully developed follicular antrum, separates the oocyte with a distinct and denser layer of granulosa cells, called cumulus cells. Mammalian oocytes remain arrested at prophase I until

LH levels surge at puberty [27]. LH causes terminal growth changes in both the oocyte and granulosa cells, resulting in the expulsion of the oocyte-cumulus cell complex from the follicle during ovulation. Additionally, LH causes the remaining granulosa cells undergo luteinisation to become the corpus luteum after ovulation, which provides endocrine support for the pregnancy [25].

Oocytes arrested at prophase I have intact nuclear envelopes and are referred to being at the germinal vesicle (GV) stage [Figure 1]. A few hours after the LH surge, the nuclear membrane of the germinal vesicle breaks down (GVBD). GVBD is a visible marker that meiosis resumption has occurred. Also at this time, the dictyate chromosomes complete prophase I and arrange themselves on the first meiotic spindle. Meanwhile, cytoplasmic contact ceases between oocyte and cumulus cells [25], causing a decrease in cyclic adenosine monophosphate (cAMP) levels in oocytes and triggering meiotic resumption [28]. The activity of maturation promoting factor (MPF), consisting of Cyclin B1 and CDK1, is required for the oocyte to resume meiosis and MPF becomes inactivated once the oocyte completes meiosis. With GVBD, chromosomes start to condense, tubulin becomes polymerized and causes homologous chromosomes to congregate on the metaphase I spindle [23]. Completion of meiosis I will cause the extrusion of the first polar body. After meiosis I completion, oocytes directly continue onto meiosis II without an intervening round of DNA replication and arrest at metaphase of meiosis II (MII). Cytostatic factor (CSF) is meiosis-specific as it maintains the oocyte arrested at MII [28]. The process of development of the oocyte, undergoing the meiotic cycle until the metaphase II stage, is known as oocyte maturation [29].

Fertilization is what triggers the second resumption and completion of meiosis regulated by oscillations of the protein kinase, MPF. This can also be triggered by parthenogenetic activation which is the induction of the initiation of the embryonic development in the absence of sperm and can be elicited by exposing oocytes to calcium ionophores, alcohol or hyaluronidase (an enzyme present in sperm required to dissociate the cumulus cell matrix). A fertilized egg is also referred to as a 1-cell embryo or a zygote. In early zygotes, collected 21 hours after hCG injections, there is resumption of meiosis by the female set of chromosomes and there is extrusion of the second polar body [23]. Moreover, the decondensation of the sperm nucleus occurs and the formation of the haploid male pronucleus takes place. By the late zygote stage, collected 26 hours after hCG injection, the nuclear membrane is formed around both the male and female haploid chromosomes producing the female pronucleus [23]. Within 6 hours

6 after fertilization, the pronuclei migrate toward the centre of the zygote, they duplicate their DNA, and their membranes break down and chromosomes assemble on the first mitotic spindle forms. This process of when the male and female haploid pronuclei approximate one another and their nuclear membranes interdigitate is known as syngamy [Figure 2] [30]. After the formation of the mitotic spindle, chromosomes congress prior to first mitotic division and a cleavage furrow develops. Following the zygote stage is a series of cleavage stages, as in the 2- cell, 4-cell, 8-cell embryo, 8-16-cell compacted morula, and early and late blastocyst stages.

GV Arrest/ GVBD Metaphase Anaphase PB Metaphase Early Zygote Late Zygote Prophase I I I Extrusion I II Ooplasm ♀ ♂ ♂

Nucleus No nuclear Meiotic Homologous Sister Zona Pellucida PB PB II Pronuclear envelope spindle chromosomes chromatids Sperm Cumulus Formation cells Figure 1: Oogenesis. GV – Germinal vesicle; GVBD – Germinal vesicle break down; PB – polar body. Detailed description of oogenesis is found in text.

1.4 EARLY MAMMALIAN EMBRYONIC DEVELOPMENT At about 12 hours prior to ovulation, during meiotic maturation, mouse oocytes become transcriptionally quiescent, while translation of the mRNA pool continues throughout the final stages of meiosis [26]. Upon fertilization, the mature oocyte is triggered to complete meiosis and yields the formation of a zygote. A minor activation of transcription occurs at the pronuclear stage of zygotes and then embryonic transcription increases at the two-cell embryo stage [30].

During preimplantation development, there is a succession of cleavage divisions from the zygote, then the two-cell, four-cell, eight-cell embryos, followed by the morula and finally the blastocyst. The blastocyst then undergoes zona hatching or lysis of the zona pellucida allowing for uterine wall implantation of the blastocyst. The progression up until blastocyst implantation in mice spans 3 days, while in humans takes 4.5 days.

Preimplantation embryonic development is characterized by a series of defined periods of gene expression, including: degradation of maternal transcripts, activation of the zygotic genome, activation of the mid-preimplantation genome, morula compaction, and blastocyst cavitation [31]. These defined periods of gene expression are present to regulate the progression

7 of development from maternal to embryonic control. Firstly, maternal products such as RNA and protein accumulate during oogenesis and are required for cleavage-stage development [Figure 2], prior to embryonic control of development. The first phase of events in the transition from maternal to embryonic control is the degradation of maternal mRNA. By the two-cell embryo, there is about 25% of remnant maternal poly-adenylated RNA and about 50% of remnant maternal protein [30].

Following the degradation phase is the zygotic genome activation (ZGA) phase, also referred to as the embryonic genome activation phase or the maternal-to-zygotic transition and this is the phase with the most marked genetic programming [31]. During this transition, the developmental program, which is initially controlled by maternal factors, gradually becomes under the control of zygotic factors as products from the ZGA are necessary for development beyond the 2-cell stage [32]. In mice, the ZGA begins during the late 1-cell zygote stage, is dominant during the 2- to 4-cell embryo stage and gradually phases out [31]. In humans, the ZGA occurs at the 4-8-cell embryo stage [33]. Cellular arrests, in both mouse and human embryos, occur if the switch from maternal to zygotic control does not adequately take place [32] and defects in this transition result in embryo fragmentation and demise. These developmental phases are depicted in Figure 2. Later stages of development are required for cell-lineage specification [31].

Oogenesis (Ovary) Embryogenesis (Oviduct)

Ovulation Fertilization

Primordial Germinal Mature Oocyte Fertilized egg Fertilized egg Syngamy Two-cell germ cell vesicle Embryo oocyte + Sperm Early Zygote Late Zygote

Cleavage furrow

Accumulation of Degradation of maternal RNA and Activation of the embryonic genome maternal RNA and protein protein Figure 2: Early Mammalian Embryonic Development. Oogenesis occurs in the ovary. After ovulation the mature oocyte, which is referred to as the metaphase II-arrested oocyte, gets released from the ovary and moves to the oviduct where it congresses with the sperm and becomes fertilized. Fertilization yields the zygote or the one-cell embryo with the maternal and paternal pronuclei. Syngamy is the developmental process when the parental pronuclei membranes breakdown and the haploid chromosomes congress on the first mitotic spindle. Following this is the formation of the two-cell embryo with the cleavage furrow. Detailed description is found in text.

During gametogenesis, haploid female and male genomes are methylated. After fertilization, the paternal genome is actively demethylated within hours of conception, while the maternal genome is progressively demethylated with each cell division, resulting in hypomethylated DNA in cleavage-stage embryos [30]. Interestingly, DNA methylation is what primarily prevents parthenogenetically activated eggs from progressing beyond the blastocyst stage, due to the monoallelic expression from epigenetic modifications [34].

Eukaryotic DNA is packed on an octamer of four core histone proteins, consisting of two molecules each of H2A, H2B, H3, and H4. Histones are crucial targets for regulation of DNA- mediated events, such as transcription, replication and DNA repair [35]. Centromere structure and function can be regulated by post-translational modifications of histones, such as

9 phosphorylation and methylation events [36]. After fertilization, the male genome undergoes decondensation, removal of protamines and re-packaging with maternal histones in a process that encompasses DNA breaks that require repair [37]. After each haploid genome is fully decondensed and packaged with histones, a pronucleus forms and undergoes DNA replication [30]. Histone modifications allow for flexible, but heritable reprogramming of the genome [38] and ensure appropriate gene activation during pre-implantation development [39]. For example, methylation of histone H3 at lysine 9 (H3K9me) is associated with gene repression and can trigger DNA methylation [40].

1.5 MATERNAL-EFFECT FACTORS Although the haploid sperm is necessary for providing DNA for the male pronucleus and is essential for egg activation, there is a disproportionate burden on the female gamete to endure successful initiation of development. The early embryo is almost entirely dependent on the oocyte for its initial supply of components that support proper early embryo development prior to the activation of the embryonic genome. Maternal-effect factors are maternally-produced molecules, such as mRNA and proteins, which accumulate in oocytes during follicular development and oocyte growth, and are utilized during embryogenesis. Maternal-effect factors are involved in the completion of meiosis, the initiation of mitosis, and the activation of the embryonic genome [41]. After fertilization, maternal-effect factors function to process the paternal genome, which is necessary for its participation during embryogenesis. Another responsibility of these factors is to eliminate the maternal store of these transcripts and proteins in due course. A final role of maternal-effect genes is to activate the embryonic genome at cleavage-stage development, which is required for development beyond the two-cell stage in mice [42]. Maternal-effect genes that have mutant embryonic phenotypes reflect the genotype of mother, not the offspring. A common phenotype among maternal-effect gene mutants is developmental arrest at cleavage-stage embryogenesis, indicating that embryonic gene transcription or translation is required to take place to replenish the depleted maternal components. Nlrp5 was one of the earliest identified maternal-effect factors discovered in mammals in 2000 [43], while the earliest identified human maternal-effect gene candidate was Nlrp7 [44]. Understanding the molecular pathways of maternal-effect factors can potentially shed light into the aetiology of clinical infertility and recurrent spontaneous abortions.

Table 1: Examples of Maternal-Effect Genes and their Phenotypes

Gene Knockout Phenotype Reference Nlrp5 -two-cell arrest in heterozygotes and Nlrp5-deficient females [43] are sterile TAp73 -lack of TAp73 in developing oocytes leads to a failure of [45] preimplantation embryonic development -i.e., decreased 4-8 cells, morula, blastocyst Filia -transcripts detected in growing oocytes and appeared to [42, 46] decrease in fully grown oocytes, but still detected until morula and early blastocyst stage -knockout female mice display decreased fecundity and delayed embryonic progression Floped -two- or four-cell arrest and females are infertile [47, 48] Padi6 -two-cell arrest, sterile [49] -decreased transcription, translation, absent cytoplasmic lattices Zygote arrest 1 -one-cell arrest and females are sterile [50] (Zar1) Nucleoplasmin2 -knockout female mice are subfertile or are infertile [51] (Npm2) -reduced cleavage to 2 cell stage -one-cell embryos do not have nucleoli and show no nucleolar clearing Cdc20 -Cdc20-null mice are embryonic lethal: permanent arrest at the [52] two-cell embryo stage Autophagy- -knockout embryos do not proceed beyond the 4-8 cell stage [53] related 5 (Atg5) -knockout embryos have reduced protein synthesis rates For additional Maternal-effect genes see reference [30].

11 1.6 NALP5/MATER

1.6.1 STRUCTURAL DOMAINS AND NALP FAMILY MEMBERS NALP5 (capital letters) is the mouse protein, is abbreviated for NACHT, LRR and Pyrin domains-containing protein 5, and is synonymous with maternal antigen that embryos require (MATER) and ooplasm-specific protein 1 (OP1). In mice, the NALP5 protein is encoded by the Nlrp5 (small caps, italics) gene, formerly known as Mater.

Based on protein domain structure, NALP5 belongs to the NALP group of proteins [54]. NALP protein family members are named for their structural domains consisting of: (1) a NACHT domain, predicted to bind to ATP; (2) a NACHT-associated domain (NAD); (3) a carboxyl-terminal leucine-rich repeat (LRR) region, known to mediate protein-protein or protein-carbohydrate/lipid interactions and (4) an amino-terminal pyrin domain (PYD), which via protein-protein interaction is directly involved in apoptotic and inflammatory signalling pathways. Currently, there are 14 genes identified as members of the Nlrp family in humans with limited information about their expression pattern and physiological role. Functionally, it has been shown that some NALPs, such as NALPs 1-3 create a molecular platform for the formation of protein complexes called inflammasomes. These complexes lead to the cleavage of caspase-1 or -5, process interleukin 1and Nuclear Factor B (NFB) activation in response to a variety of bacterial and viral products, thus implicating NALPs as regulators of innate immunity responses [55]. In addition, NALP family members have also been linked to the regulation of apoptosis [54]; however, the exact molecular cell death pathways, downstream of the NALP response, have not yet been determined. Recently, multiple Nlrp family members, including Nlrp5, Nlrp8 and Nlrp9, have been shown to have oocyte- and embryo-restricted expression in bovine and thus have roles implicated with female reproduction [56]. Nlrp14 in mice is required for normal preimplantation development, as observed by arrested embryos at the 1-cell and 8-cell stages following siRNA injection into zygotes [57]. Mutations in human Nlrp7 have been associated with recurrent hydatidiform moles, spontaneous abortions, still births and intrauterine growth retardation [44, 58]. In addition, Nlrp7 overexpression in humans has been related to the development of testicular seminomas, a germ cell tumour [59]. Likewise, about 3% of men with a mutation in Nlrp14 developed azoospermia or severe oligozoospermia, indicative of decreased male fertility or even sterility [60]. These examples highlight Nlrp family members as key players in reproduction.

1.6.2 IDENTIFICATION OF NALP5 NALP5 was originally identified as an oocyte-specific antigen for murine autoimmune premature ovarian failure, which was induced in mice by performing neonatal thymectomy [61]. This syndrome is similar to premature ovarian failure in humans, caused by an autoimmune response, and it triggers the onset of early menopausal symptoms and infertility in young women. Recently, however, NALP5 has been found to be an autoantigen involved in hypoparathyroidism in patients with autoimmune polyendocrine syndrome type 1 (APS-1) and in women NALP5 localizes to ovaries and parathyroid glands [62]. About 68% of female patients who have APS-1 and generate antibodies against NALP5 have hypogonadism, an ovarian dysfunction where ovaries contain very few follicles and thus produce little or no hormones [62]. These findings allude to the possibility that NALP5 protein in women may potentially be linked to premature ovarian failure.

Although Nlrp5 was one of the earliest identified maternal-effect genes in mammals in 2000, relatively little information is known about the function of the gene and protein [63]. In 1999, Tong and Nelson isolated and characterized the NALP5 protein and found that it has a molecular mass of 125 kDa and is composed of 1111 amino acids. In germinal vesicle (GV)- stage oocytes, NALP5 protein was found to localize to the ooplasm, mitochondria, nucleolus and close to nuclear pores [64]. Nlrp5 is a single-copy gene that is expressed in exclusively in oocytes in mice and is found on the proximal end of chromosome 7 in mice [63]. The mouse Nlrp5 gene locus spans about 32 kb DNA and is composed of 15 exons and 14 introns [63]. The mouse NALP5 protein has an uncharacterized amino-terminal domain, instead of having the Pyrin domain that is present in human NALP5 [54] [Figure 3]. In addition, mouse Nlrp5 is a potential gene candidate for human sterility, since human and mouse Nlrp5 share a 67% nucleotide sequence homology at the cDNA level and a 53% homology at the amino acid level [65].

Figure 3: Mouse and Human NALP5 protein structure. NACHT domain-green; Leucine Rich Repeats-blue; Pyrin domain-gray.

1.6.3 NALP5 EXPRESSION PATTERN During oogenesis in mice, Nlrp5 is transcribed and mRNA transcripts are detected from the stage when primordial oocytes make their decision to become primary oocytes [64]. Nlrp5 transcript levels peak at the germinal vesicle stage and dramatically decline by the two-cell stage, becoming undetectable in preimplantation embryos. Because Nlrp5 transcripts are degraded before the late-one-cell to two-cell embryo (period of embryonic genome activation in mice), this suggests that the NALP5 protein product is exclusively derived from the maternal genome [64]. NALP5 protein is first detected in small primary follicles and is present in all stages of preimplantation embryos up until the early blastocyst stage [64]. NALP5 protein subcellularly localizes to the mitochondria, nucleoli, and close to the nuclear pores of germinal vesicles of oocytes, as determined by immunogold electron microscopy [64].

Recently, Nlrp5 transcripts and protein have been found in bovine up until the 16-cell embryo stage [66], in non-human primates (rhesus macaque monkeys) they were present up until the 8-cell stage [67], in pigs Nlrp5 transcripts were present in germinal vesicles [68], and humans transcripts were observed at the germinal vesicle stage [65] and day-5-embryo stage [69]. Aside from the mentioned localization of human NALP5 protein in ovaries and parathyroid glands [62], a recent report found that human NALP5 localizes to oocytes, embryos [69], and also cumulus cells [70]. Furthermore, in humans NALP5 was discovered to be found in oocytes and follicle cells that are spatially and functionally associated with the oocyte suggesting that in humans NALP5 has an earlier role compared to other mammalian species [70]. The only other report of NALP5 localization aside from the mentioned regions, claimed to have found NALP5 in neuronal cultured cells that were subjected to apoptotic injury [71]. Taken together, Nlrp5 transcripts and NALP5 protein are consistently expressed in female gametes and preimplantation embryos in a vast range of mammalian species, including humans, implying that Nlrp5 and NALP5 likely have a role in female infertility.

Additionally, in microarray analysis performed by Hamatani et al., it was discovered that metaphase II oocytes from old mice (aged 42-45 week-old) exhibited a two-fold decrease in Nlrp5 transcript levels, compared to oocytes from young mice (aged 5-6 week-old) [57]. These findings suggest that the expression of Nlrp5 transcripts decreases with increased maternal aging of oocytes.

1.6.4 NLRP5-DEFICIENT PHENOTYPE The Nlrp5-(homozygous) deficient, also known as knockout, mouse model was created using homologous recombination in embryonic stem cells to mutate the second Nlrp5 intron without interrupting the adjacent exons, by insertion of a neomycin cassette [42, 64]. By northern blot analysis, although greatly diminished compared to wildtype and Nlrp5 heterozygous eggs, Nlrp5 mRNA was still detectable in the Nlrp5 knockouts [42]. Likewise, by western blot analysis, they also found that there were residual levels of NALP5 protein present in Nlrp5 knockouts, suggesting that the knockout mutation destabilizes the Nlrp5 transcript and causes a severe hypomorph phenotype, affecting early mouse development [42]. In this thesis, Nlrp5 knockout females, oocytes, or embryos are also referred to as being Nlrp5-deficient.

Nlrp5 knockout male mice have no known phenotypic abnormalities. Female knockouts undergo normal oogenesis, ovarian development, oocyte maturation, ovulation and fertilization [64]. Additionally, Nlrp5-deficient female mice respond normally to exogenous gonadotropin stimulation seeing as they release oocytes of comparable number and morphology to wild-type females [64]. Wildtype zygotes developed to the 2-cell embryo, morula and blastocyst stages. However, zygotes originating from Nlrp5-deficient oocytes, with either the Nlrp5 knockout or Nlrp5 heterozygote genotype, remained arrested at the 2-cell stage and degenerated, while only a small fraction reached the blastocyst stage [43]. As a result, Nlrp5-deficient females are sterile. This developmental arrest of embryos at the 2-cell stage cannot be rescued by the fertilization with wildtype sperm [43]. Thus, Nlrp5 is a maternal-effect gene that is required for embryos to progress in development beyond the 2-cell stage.

Further evidence from the knockout mouse model showed that de novo RNA transcription was considerably reduced in Nlrp5 knockout one- and two-cell embryos [43]. Two-cell embryos lacking Nlrp5 were able to synthesize 60% of the normal levels of the transcription-related complex, which is a marker for embryonic genome activation [43]. Although Nlrp5 may not be crucial for initiation of all transcription-translation machinery in early embryos, it is necessary for transcription in early embryos. Moreover, though NALP family members are structurally related, as mentioned earlier, there seems to be no functional redundancy in Nlrp5-deficient mice since the absence of Nlrp5 alone causes infertility [43].

The Nlrp5 knockout mouse model is useful means to study human infertility since mouse and human Nlrp5 genes and proteins are conserved [65]. The study of Nlrp5 pertaining to

15 primate fertility is further supported by work done in rhesus macaque primates, confirming that NALP5 localizes to oocytes [67]. Furthermore, especially since Nlrp5 is exclusively present in oocytes, research in the field of oocyte development is limited by human samples of oocytes. For instance, it is difficult to get healthy samples of human oocytes and embryos, since the primary source of human gametes and embryos is from subfertile couples because they are the ones who seek assisted reproductive technology treatment and are most likely to donate their eggs for research [72]. In suboptimal quality oocytes, Nlrp5 mRNA or protein levels may already be altered (i.e. may be deficient/excessive) and we would not have a suitable comparison with control group of healthy human oocytes.

1.6.5 NALP5 INTERACTING PARTNERS With all available literature to date, the function of NALP5 remains elusive, although only recently some light has been shed on the interacting members of this protein. NALP5 has been discovered to be one of the four proteins that contribute to the Subcortical Maternal Complex [47]. This complex assembles during oocyte growth and is fundamental for zygotes to develop beyond the first embryonic cell division. The subcortical maternal complex is composed of physical interactions between: NALP5, FLOPED, TLE6 and FILIA and localizes to the subcortex of oocytes and preimplantation embryos [Figure 4]. Absence of this complex drastically impedes development beyond the two-cell stage [47]. This phenotype has been suggested to be caused by abnormalities in syngamy, mitotic spindle formation, cytokinesis or cell cycle progression [47].

In the Floped (factor located in oocytes permitting embryonic development)-null mouse model, knockout females have normal ovarian physiology, however less than 20% of Floped- deficient embryos develop beyond the two-cell stage [47], thus resembling the Nlrp5 phenotype.

TLE6 (transducin-like enhancer of split 6) is a putative transcriptional co-repressor and it has a developmental transcript profile which is similar to that of Nlrp5 in that transcripts accumulate during oogenesis, peak at the germinal vesicle stage and degrade drastically at the two-cell stage [47]. TLE6 has been shown to contain a nuclear localization signal [30] and Tle6 is a homologue of the Groucho gene in Drosophila, which is an inhibitor of dorsal (NFB) signalling. Nonetheless, the function of Tle6 in oocytes and embryos is currently unknown.

Plasma Membrane TLE6

FLOPED

NALP5

FILIA

Figure 4: Subcortical Maternal Complex. The subcortical maternal complex localizes proximal to the plasma membrane in oocytes and preimplantation cleavage-stage embryos. NALP5 interacts with all of the proteins belonging to the complex, including TLE6, FLOPED and FILIA.

By comparing protein profiles of ovulated eggs from wildtype and Nlrp5 knockout mice, FILIA was identified as a binding protein of NALP5 [42]. Nlrp5-deficient oocytes have a dramatic decrease in the abundance of FILIA protein. Filia has two isoforms, as detected by northern blot analysis. Ohsugi et al. also showed that the near absence of Nlrp5 mRNA did not affect Filia mRNA expression; however, the decrease in NALP5 protein caused a profound decrease in FILIA expression, thus the stability of FILIA is dependent on the presence of NALP5 [42]. Filia knockout female mice are viable and are fertile although they have reduced fecundity, as evidenced by smaller litters [46]. The absence of Filia does not disturb NALP5, but it does cause chromosomal abnormalities in embryos [46]. Filia-deficient female mice showed an increased rate of aneuploidy during the 2-cell stage supported by increased frequency of lagging chromosomes, micronuclei, and hyperploidy. They also found that Filia knockouts had increased abnormalities in mitotic spindles due to altered allocation of certain Spindle Assembly Checkpoint proteins [46]. These findings suggest that Filia plays a role in ensuring chromosome stability and euploidy during mouse embryogenesis.

1.7 SPINDLE ASSEMBLY CHECKPOINT The meiotic and mitotic spindles are the cellular structures that are responsible for proper chromosome segregation in all cell types. The meiotic spindle’s role is critical in germ cells, including oocytes, since inadequate chromosomal segregation in gametes may lead to aneuploidies, causing unexplained infertility, miscarriages and congenital birth defects [73]. Spindle microtubules are hollow, cylindrical polymers that are assembled from α and β-tubulin heterodimers [74] and form barrel-shaped spindles that can be visualized by immunocytochemistry with an anti-α-tubulin antibody [Figure 5B]. During development in the M-phase of cell cycle an essential process occurs that maintains genomic integrity by segregating chromosomes faithfully. Chromosomes congregate on the equatorial plate of bipolar spindles for segregation, cross-linking their kinetochores to the emanating microtubule arrays [Figure 5]. Kinetochores are protein structures that localize to the centromeric region of sister chromatids [75].

Equatorial plate/ metaphase plate

Chromosomes Microtubules α –tubulin

Pericentriolar material

Kinetochores A. B.

Figure 5: Metaphase II-arrested meiotic spindles. A. Schematic representation of a metaphase II-arrested (MII) meiotic spindle. B. Immunocytochemistry and confocal imaging of a mouse MII spindle stained with α-tubulin (green). Chromosomes aligned at the metaphase plate are visualised with DAPI (blue).

The Spindle Assembly Checkpoint (SAC) is a regulatory mechanism that prevents errors in chromosome segregation and aneuploidy during both mitosis and meiosis [76]. The SAC is a complex of proteins that monitors kinetochore-microtubule attachment and tension. Also, this complex functions to prevent anaphase onset until all kinetochores are properly aligned to bipolar spindles at the metaphase plate [77]. In mitosis, even a single unattached kinetochore to microtubule causes the recruitment of components of the checkpoint, initiating a signalling cascade that results in CDC20-dependent inhibition of the anaphase promoting complex or

18 cyclosome (APC/C), hence delaying anaphase onset and pausing meiosis progression [78, 79]. The APC/C is an E3 ubiquitin ligase that drives the transition between metaphase to anaphase, by catalyzing the proteasomal degradation of anaphase inhibitors and mitotic or meiotic cyclins, such as securin and cyclin B1. More specifically, when there is an unattached kinetochore to microtubule, SAC proteins form a complex and bind to and inhibit CDC20 [Figure 6A]. This inhibition of CDC20 occurs by phosphorylation events at four phosphorylation sites in Xenopus, consequently causing the inhibition of the APC/C and causing a pause in the cell cycle [80] until the defect gets corrected, at which point cell cycle will continue.

BUB SAC complex 3 TAp73 Cell cycle is forms on BUB BUB APC/C paused at unattached MAD1 R1 1 metaphase K-MT CDC20 MAD 2

SAC complex Cell cycle does not form CDC20 APC/C continues with on attached anaphase onset K-MT

Figure 6: Spindle Assembly Checkpoint mode of action. Several proteins form the SAC complex and bind to and inhibit CDC20, an activator of the anaphase promoting complex (APC/C). These proteins include: BUB1, BUB3, BUBR1, MAD1, MAD2 and TAp73. A. When there is an unattached kinetochore to a microtubule (K-MT), these proteins bind to and phosphorylate CDC20 causing its inhibition. This in turn inhibits the APC/C, causing SAC activation and metaphase arrest, preventing anaphase entry until proper K-MT attachment is attained. B. When there is proper kinetochore to microtubule attachment; the SAC complex of proteins does not bind to CDC20. CDC20 activates the APC/C, causing anaphase entry.

When all kinetochores have properly attached to microtubules with the correct tension by pulling forces, the SAC is silenced since CDC20 is not inhibited by the SAC complex of proteins, thus causing proper APC/C activation [Figure 6B]. This in turn, allows for anaphase resumption and cell cycle progression. The molecular pathway of the SAC has been highly conserved throughout eukaryotic evolution [81]. Additionally, the molecular activation of pathways triggering the SAC has been intensively studied in mitosis of somatic cells; however its relevance and conservation of the pathway in meiosis is currently not well understood. The

SAC players that will be investigated in detail are: BUB1, BUBR1, TAp73, Aurora kinases B and C, and Dynactin p50. More SAC proteins are listed and described in Table 2. Table 3 highlights some of the frequency of abnormalities observed with spindle assembly checkpoint deficiencies in mutant mammalian models.

Chromosome Segregation during Meiosis of the female germ line

In meiosis I homologous chromosomes segregate, while sister chromatids remain intact. Sister chromatids are held together along the entire chromosome arm and centromere by a cohesin complex of proteins containing Rec8 [28]. Polo-like kinase 1 (Plk1) phosphorylates Rec8 at chromatid arms, which causes the degradation of the cohesin complex by separase. Meanwhile, BUB1 recruits a complex of Shugoshin and phosphatase PP2A to centromeres to protect Rec8 from phosphorylation by Plk1 and thus its degradation [82]. Separase cleaves Rec8 to release cohesin between sister chromatid arms, while centromeric cohesins are kept unphosphorylated by PP2A. This causes separation of the sister chromosomes facilitating the onset of anaphase of meiosis I. Cyclin B and securin get degraded after ubiquitination by the APC/C.

Meiosis II entails the segregation of sister chromatids. The cytostatic factor component, Emi2 (early mitotic inhibitor 2), inactivates the APC/C during metaphase II [28]. Emi2 is responsible for the arrest of oocytes at metaphase II before fertilization. Degradation of Emi2, at fertilization onset, activates the APC/C, causing cyclin B and securin to get degraded. Separase then cleaves Rec8 at the centromeres of sister chromatids for separation during anaphase II [28].

Serine/threonine protein kinase BUB1 (Budding Uninhibited by Benzimidazole), also known as an “attachment sensor,” is a core component of the spindle assembly checkpoint complex that localizes to kinetochores that are unattached to microtubules [83, 84]. In somatic cells, BUB1 dissociates upon proper kinetochore to spindle attachment and its protein expression decreases [85]. BUB1 has been shown to be a master regulator required for the assembly of multiple checkpoint proteins at the kinetochore [85], specifically, BUB1 is required for the localization of BUBR1 and MAD2 to the kinetochore. BUB1 has been shown to bind to BUBR1, MAD1, MAD2 and BUB3 [77]. In somatic cells, BUB1 is recruited to kinetochores that are unattached to microtubules, early in prophase of mitosis. Loss of BUB1 in Drosophila

20 causes chromosome missegregation and lethality [86]. It was recently reported that Bub1 knockout mice are embryonic lethal and that Bub1 heterozygous mouse oocytes were aneuploid, due to premature sister chromatid separation, and exhibited decreased fecundity [87]. In wildtype mouse oocytes, BUB1 is phosphorylated and remains associated with kinetochores during the metaphase arrest of meiosis II, indicating that this protein is involved in the SAC in mammalian oocytes [88].

BUBR1, also known as the vertebrate homologue of the MAD3 (mitotic-arrest deficient 3) protein, is a SAC protein kinase with a defined role in mitosis. Also, it has recently been identified as a SAC protein involved in meiosis progression [89]. BUBR1 becomes recruited to the kinetochores, after BUB1 localization during late prophase [85]. BUBR1 forms a stable complex with CDC20 and causes its inhibition via a phosphorylation event [80]. In mice, BubR1 deficiency causes mouse embryonic fibroblasts to undergo premature sister chromatid separation, indicative of disrupted SAC function [90]. Moreover, a decrease in BubR1 (hypomorphic allele) causes oocytes to have abnormal metaphase II configuration, resembling aged oocytes, suggesting that this observed chromosomal alignment phenotype in these mice is what leads to the observed infertility and early aging [90].

Another protein of interest is p73, encoded by the Trp73 gene. p73 is a transcription factor which plays a role in the regulation of development, cell death and cell fate commitment [45]. The Trp73 gene has two promoters that drive the expression of two isoforms, an amino- truncated (DeltaNp73) isoform and a transactivation domain-containing (TAp73) isoform, each with various splicing isoforms. Our laboratory has previously analysed the role of the transactivation domain-containing isoform of p73, TAp73, in oocytes and found that TAp73 is a novel maternal-effect gene whose deficiency leads to a failure of preimplantation embryonic development and causes infertility [45]. TAp73-deficient females ovulate fewer oocytes and those that are produced have higher rates of spindle abnormalities and multinucleated blastomeres. Additionally, fewer zygotes, originating from TAp73-null females, progressed to the blastocyst stage. Recently, it has been found that TAp73 regulates SAC functions in both mitosis and meiosis, by modulating BUBR1 activity [91]. In the absence of TAp73, metaphase II-arrested oocytes exhibit increased rates of spindle defects, misalignment and unattached

21 chromosomes, likely due to insufficient recruitment of BUB1 and BUBR1 proteins to kinetochores.

The Aurora kinases are family of serine/threonine kinases that are required for appropriate chromosomal segregation during mitosis or meiosis [92]. Moreover, Aurora kinases are involved in regulating chromatin status via phosphorylation of histones [Figure 7]. Aurora kinase B and Aurora kinase C share a high sequence identity of 78% [93]. Aurora kinase B is a chromosomal passenger protein that is essential for proper chromosome alignment, chromosome segregation and cytokinesis [94, 95]. In somatic cells, after chromosome biorientation is complete, activation of the anaphase promoting complex causes the removal of Aurora B from the centromeres and allows cells to progress to anaphase [96]. Furthermore, Aurora B has been found to modulate chromosomal alignment in mouse oocytes [92]. It has previously been shown that by microinjecting an Aurora B-tagged GFP protein into germinal vesicle-stage oocytes, this kinase is enriched on the centromeres or kinetochores [92]. Previous reports attempting to localise endogenous Aurora B by immunocytochemistry in metaphase II mouse oocytes found that there was low to undetectable levels of endogenous Aurora B protein [92, 97]. Aurora kinase C is the germ cell-specific Aurora kinase that is found in oocytes and spermatocytes [98]. Moreover, it has recently been shown that in mouse oocytes, Aurora C deficiency causes chromosomal misalignment, abnormal kinetochore-microtubule attachment, premature chromosome separation and cytokinesis failure in meiosis I and causes the production of large polyploid oocytes [97]. Knocking down Aurora C mRNA in prometaphase I oocytes causes inhibition of BUB1 and BUBR1 localization [97]. It has been reported that Aurora C localizes to the centromeres in metaphase II oocytes [97].

cdc2 Thr288

Aurora B & P Aurora C Ser10

Histone P H3

Figure 7: Aurora kinase pathway. Phosphorylation of Aurora kinases, B and C, triggers the phosphorylation of histone H3.

Pan-Aurora kinase inhibitor (ZM447439) studies in mouse oocytes show that there is failed meiotic progression to metaphase II, chromosomes are improperly condensed and

22 chromosomal alignment is perturbed [92, 99]. These findings suggest that Aurora kinases function in promoting the fidelity of chromosome segregation during meiosis in oocytes.

Dynactin p50 is also referred to as dynamitin and dynactin subunit 2 and is a microtubule protein that is involved in the intracellular, bidirectional transport of organelles by binding to and activating the dynein motor protein [100]. Dyanctin p50 is also needed for the release of the MAD1, MAD2, and BUBR1 complex from aligned kinetochores [77]. In somatic cells, during metaphase of mitosis, dynactin p50 disappears from the kinetochores upon chromosomal alignment at the metaphase plate, but has increased localization at the kinetochores of nonaligned metaphase chromosomes [101].

Table 2: List of Spindle Assembly Checkpoint Proteins and their Phenotypes

Protein Localization Molecular Phenotype References BUB1 -kinetochores -Protein kinase that phosphorylates Cdc20 in response to [78, 85, 88] -binds to improper kinetochore-microtubule attachment, which BUB3, inhibits APC/C BUBR1, -in meiosis, helps recruit shugoshins to kinetochores and to MAD1, MAD2 maintain Rec8 at sister kinetochores until meiotic anaphase II onset BUBR1 -kinetochores -recruits Mad1 and Mad2 to kinetochores lacking [91, 102] -forms complex attachment and tension with CDC20, -protein kinase that controls the activation of APC by MAD1, MAD2, binding and inhibiting p55cdc20 BUB3 BUB3 -kinetochores -in cases of unattached kinetochore-microtubule, helps [103] -binds to BUB1 inhibit the APC BUBR1 MAD1 -kinetochores -phosphoprotein that recruits MAD2 to kinetochores [28] -binds to lacking attachment and tension MAD2, CDC20, BUBR1 MAD2 -kinetochores -in cases of unattached kinetochore-microtubule, MAD2 [104] -forms complex forms complex and inhibits the APCCdc20 with MAD1, CDC20, BUB1 BUBR1 CDC20 -forms complex -recruits substrates to the APC/C [78, 105] (Cell with MAD2, -is a co-activator of the APC/C Division BUBR1, BUB3 Cycle) APC/C -activated by -Multi-subunit E3 ubiquitin ligase [78, 102] (Anaphase- CDC20 -APCCdc20 controls the degradation of cohesive ties between Promoting chromosomes (such as securin, cyclin B, Shugoshin1) at the Complex) metaphase-anaphase transition

Table 2: List of Spindle Assembly Checkpoint Proteins and their Phenotypes (Continued)

Protein Localization Molecular Phenotype References BRCA1 -spindle poles -in mouse oocytes, BRCA1 regulates meiotic spindle [106] assembly and the spindle assembly checkpoint TAp73 -spindles -modulates BubR1 activity [45, 91] -directly interacts -BUBR1-p55Cdc20 interaction was impaired in TAp73- with BUB1, deficient cells BUBR1, BUB3 Shugoshin -centromeres -meiosis-specific SAC protein that keeps sister chromatids [107] 1 together during meiosis I, by protecting Rec8 from separase activity during meiosis I and not meiosis II Rec8 -centromeres -meiosis-specific SAC protein; centromeric cohesion [28] protein that is cleaved by separase at anaphase onset Polo-like -spindle poles -phosphorylates meiotic cohesion proteins targeting them [28] Kinase 1 for proteolytic cleavage by separase (in metaphase I and II) Aurora A -centrosomes -critical microtubule organizing centre (MTOC)-associated [108, 109] kinase component involved in the continuation of meiosis, MTOC multiplication, correct spindle formation Aurora B -kinetochores -member of chromosome passenger complex [92, 96] kinase -APC activation causes the removal of Aurora B from centromeres Aurora C -centromeres -germ cell-specific kinase [97] kinase -important for biorientation of chromosomes in meiosis I

Table 3: Frequency of Spindle Configuration and Chromosomal Segregation Defects

Mutated Gene Frequency of Abnormality References Filia Improper mitotic spindles in 2-cell embryos: KO-69%; WT-11% [46] (Knockout) Chromosome misalignment in 2-cell embryos: KO-64%; WT-11% -causing subfertility (decreased little size) TAp73 Spindle abnormalities in MII mouse oocytes: [45] (Knockout) TAp73-null-25%; WT-7% Bub1 Aneuploidy in MII oocytes:Bub1+/m-76%; WT-4% [87] (Heterozygosity) Aneuploidy in zygotes:Bub1+/m-81%; WT-10% BubR1 Premature sister chromatid separation [90] (Hypomorph) -Bub1b–/H and Bub1bH/H-15-38%; WT-1-3%

BubR1 (RNAi) Aneuploidy with 50% decreased BUBR1 expression [110] BRCA1 Spindle abnormalities in mouse oocytes [106] (RNA interference) -BRCA1 siRNA-64%; control-siRNA-11% Misaligned chromosomes [16] -BRCA1 siRNA-45%; control-siRNA-5%

1.8 DNA DAMAGE Maintenance of genome integrity is crucial to the survival and fertility of organisms. In the presence of DNA damage, cells respond by activation of surveillance mechanisms that lead to DNA repair. Thus far, little is known about DNA repair in oocytes. What is known about DNA damage/repair occurs during the early stages of meiosis, at prophase I entry. During meiotic prophase, chromosomal recombination is initiated and this promotes the formation of DNA breaks and physical links between homologous chromosomes to ensure the exchange of genetic information. In mammals, recombination initiates with DNA double-strand breaks [111], which have to be repaired by ATM kinase-mediated mechanisms [112].

Our work is focused on the ovulated oocyte, which is the final product of meiosis. A recent study found that ovulated oocytes from young mice were more capable of repairing damaged DNA relative to oocytes from old mice [113]. In oocytes, the age-dependent efficacy of DNA repair mechanisms has been linked to increased rates of aneuploidy [16].

Fully grown oocytes isolated from adult mice have previously been reported to be capable of excision repair triggered by ultraviolet (UV) irradiation [114]. This was observed by unscheduled DNA synthesis, by means of radioactive thymidine incorporation, after oocytes (GV-MII stage) were exposed to UV radiation. UV irradiation causes damaged nucleotide bases and disrupts base pairing, which are excised and repaired by oocytes [115]. UV type C is one form of UV irradiation, which is elicited at 254 nm and produces cyclobutane pyrimidine dimers [116]. The mode of UV repair is known as nucleotide-excision repair, where a small region of UV-induced DNA damage: 1) is recognized, 2) is removed by the incision of the DNA strand and excision of damaged bases and 3) undergoes repair synthesis by refilling DNA and rejoining by ligation [116]. The DNA damage signal in the nucleotide-excision repair pathway is primarily recognised by ATR (ataxia telangiectasia and RAD3-related) and causes the phosphorylation of H2AX in a CHK1 (checkpoint protein-1)-dependent manner in response to replication stress [116]. Furthermore, 53BP1 is also recruited to sites of replication stress by the ATR target, CHK1 kinase. The nucleotide-excision repair pathway is dependent on p53 transactivation, as the loss of p53 in human fibroblasts results in the reduced repair of UV- induced DNA damage [117]. In addition, the excision repair pathway is relevant to normal embryo development and it has been recently shown to occur in zygotes, as a means of repairing DNA damage, especially in the paternal pronucleus [118].

The molecular mechanism of the DNA damage response has been extensively studied in somatic cells and involves a series of proteins that actively become recruited to the sites of DNA damage. The pathways are outlined below [Figure 8] and will be further discussed in detail in the text.

Ionizing UV Damage ROS Radiation

Nucleotide/Base Double Strand DNA Excision Repair Breaks

ATM ATR Aurora B

p p ATM CHK1 p H3

p p p CHK2 H2AX 53BP1 p H2AX 53BP1

BRCA1

Figure 8: DNA Damage/Repair Response Pathway in Somatic cells. The various forms of DNA damage elicit slightly different DNA damage response pathways, which sometimes use the same machinery.

Histone H2AX, a variant of histone H2A, is required in mice to maintain genome stability [119]. H2AX knockout mice display increased levels of chromosome instability and defects in DNA repair, by means of improper recruitment and assembly of DNA repair factors, 53BP1 and BRCA1, to DNA double stand breaks [120, 121]. Phosphorylated histone H2AX (γH2AX) at serine 139 is a classical marker of DNA damage that is typically present when there are double strand breaks, but is also recruited to DNA sites in response to UV irradiation [122]. Although the role of γH2AX in response to ionizing radiation has been studied, its role in response to UV- induced DNA damage is not well understood. A recent report of UV exposure on fibroblast cells demonstrated that UV irradiation causes pan-nuclear staining of γH2AX and this is likely a reflection of the nucleotide-excision repair pathway, even though until now γH2AX has been

28 used as a marker of DNA double stranded break formation [122]. Furthermore, H2AX phosphorylation is dependent on the process of repairing the UV-elicited DNA damage, which in this case requires nucleotide excision repair factors and is dependent on the ATR pathway [Figure 8] [122]. These findings indicate that γH2AX after UV irradiation could protect cells from genome instability by maintaining the integrity of cell cycle checkpoints [122]. Additionally, the distribution of γH2AX has been investigated in preimplantation embryos in the absence of DNA damage [35]. Without DNA damage, γH2AX was detected shortly after fertilization at the zygote stage with higher accumulation on the paternal genome. This could suggest that there is an intense repair process in the paternal pronucleus compared to that of the female. Moreover, γH2AX accumulation has been shown to increase when irradiated sperm is used to fertilize the egg compared to non-irradiated sperm [37]. In normal development, γH2AX foci form, however if the paternal DNA has been damaged, the size of the γH2AX foci is further increased, suggesting that the maternal factors in the cytoplasm are sensing and/or repairing the paternal DNA.

The next DNA damage response factor that we investigated was 53BP1 (p53-Binding Protein-1), which is involved in DNA damage-induced, cell-cycle arrest. 53BP1-null mice result in growth retardation and are prone to develop tumours [123]. 53BP1 displays cytoplasmic and nuclear localization and stimulates p53-mediated transcription [35]. Upon elicitation of ionizing radiation, H2AX becomes phosphorylated (γH2AX), enabling the recruitment of phosphorylated 53BP1 to double-stranded break foci, in an ATM-dependent manner [Figure 8] [121]. With UV- induced DNA damage, the ATR pathway is elicited and H2AX becomes phosphorylated (γH2AX), allowing for the recruitment of 53BP1 in response to replication stress [Figure 8] [116]. In HeLa cells, it has previously been shown that UV irradiation (at a dose of 50 J/m2) elicits the formation of 53BP1 nuclear foci [124]. However, this may be cell type- and dose- specific, as some recent data in human fibroblasts show that 53BP1 is evenly distributed throughout the nucleus in response to 20 J/m2 of UV irradiation [122]. Furthermore, the distribution of 53BP1 was also evaluated in preimplantation embryos in the absence of DNA damage and was detected from the two-cell stage onward and was absent from mitotic chromosomes [35].

Phosphorylation of histone H3 at serine 10 (H3S10) is strongly correlated with chromosome condensation during mitosis and meiosis [125]. In addition, H3 phosphorylation is also associated with transcriptional activation during the S phase of cell cycle [95]. Recently,

29 histone H3 de-phosphorylation has been observed in somatic cells in response to variety of DNA damaging agents, including ionizing irradiation and oxidative stress, both of which cause double strand DNA breaks [95]. It was proposed that DNA damage causing double strand breaks cause the inhibition of Aurora kinase B, which then inhibits phosphorylation of its target, histone H3 [95].

BRCA1 has been implicated in cell cycle progression, DNA damage signalling and repair, maintenance of genome integrity, ubiquitination and regulation of transcription [126]. Thus, it is not surprising that the disruption of BRCA1 function causes early embryonic lethality due to massive apoptosis in embryos [127]. Also, Brca1 mutations or silencing contribute to breast and ovarian cancers. Particularly relevant to our studies, BRCA1 has been shown to be needed for meiotic spindle assembly and SAC activation during in vitro maturation of mouse oocytes [106]. BRCA1 appears to localize to spindle poles in MII oocytes and its depletion, by silencing, causes misaligned chromosomes and impaired spindles [16, 106]. Furthermore, in old mouse oocytes BRCA1 protein and transcript levels were found to decrease and higher incidences of aneuploidy were observed when Brca1 was silenced [16].

The DNA-damage signal from DNA double strand breaks is mainly recognized by ATM (ataxia telangiectasia mutated) [116]. ATM, a serine/threonine kinase that becomes activated in response to DNA damage, causes the phosphorylation of key proteins in several signalling pathways through its substrate checkpoint protein-2, CHK2. The phosphorylation status of ATM protein kinase (S1981) is a site known to be activated in response to changes in chromatin structure and this active form becomes recruited to sites of DNA double strand breaks [116]. Relevant to oogenesis, Atm knockout mice are sterile due to meiotic arrest, as evidenced by ovaries that are devoid of primary oocytes and follicles 11 days after birth [112]. This oocyte death in Atm knockouts has been correlated with a response to unrepaired or incorrectly repaired double-strand breaks in meiotic recombination [111].

1.9 RATIONALE Nlrp5 is a maternal-effect gene that is necessary for murine embryos to progress beyond the two-cell stage of development [43] and lack of the gene causes female sterility in mice. The human NALP5 and mouse Nlrp5 genes and proteins are conserved, sharing a 67% homology at the cDNA level and a 53% homology at the amino acid level, respectively [65]. Investigating a mouse model with disruption of Nlrp5 may help elucidate the molecular causes triggering the embryonic phenotypes and could provide the basis for exploring clinical implications of infertility in women.

Preliminary data from our laboratory demonstrate that a significant proportion of Nlrp5- deficient preimplantation embryos contain multinucleated blastomeres. Furthermore, cases of human embryo multinucleation are prevalent in in vitro fertilization (IVF) clinics. The transferring of multinucleated blastomeres to IVF patients has been shown to result in significant reductions in the rates of embryo implantation, clinical pregnancy and live births [21].

Along with the reasoning that the absence of Nlrp5 in embryos yields an increased incidence of multinucleated blastomeres and because the lack of Filia (NALP5-interacting partner), causes subfertility and aneuploidy due to dysregulation of spindle assembly regulators in mice, this could indicate that Nlrp5 may play a role in the regulation of the SAC and/or DNA integrity. Spindle defects and DNA damage trigger independent pathways and have checkpoints which are regulated by different molecular targets. Currently, it is not known what triggers DNA damage in ovulated oocytes and there are no established molecular markers that could reliably be used for DNA damage repair analysis in this model. In early development, the DNA damage repair response is of maternal origin since up until the embryonic genome activation, the zygote is dependent on the maternal transcripts and factors of oocytes for all developmental processes. As a result, the stored maternal transcripts are likely responsible for controlling the integrity of the chromatin [128]. Nlrp5 is a maternal-effect gene that has been proposed to regulate the transition from oocyte to embryo, as evidenced by the two-cell arrest phenotype and small changes in the protein profile and active RNA transcription in Nlrp5-deficient embryos [43]. However the role of Nlrp5 in the maintenance of DNA integrity has not been explored.

HYPOTHESES

We hypothesize that Nlrp5 plays a role in facilitating the activation of the Spindle Assembly Checkpoint (SAC) in metaphase II oocytes. We propose to investigate the role of Nlrp5 as a factor that facilitates the proper functioning of the SAC during meiosis II. In addition, we believe that Nlrp5 could play a role in triggering an appropriate DNA damage repair response in oocytes.

To address these hypotheses, we set the following objectives:

OBJECTIVES: 1. To ascertain the localization of NALP5 in metaphase II-arrested oocytes via confocal microscopy 2. To investigate markers of spindle assembly checkpoint integrity in Nlrp5-deficient oocytes 3. To determine the distribution of DNA damage repair markers in Nlrp5-deficient metaphase II-arrested oocytes

2 METHODS

2.1 MOUSE HUSBANDRY ICR mice (Harlan, Indianapolis, IN, USA) were reared at the Mount Sinai Hospital Animal Facility. The Hospital Animal Care Committee approved all animal protocols and protocols were in compliance with standards for the ethical treatment of animals. All ICR mice were housed at Mount Sinai Hospital animal facility with access to ample water and food, and were kept in a 12:12 h light: dark cycle.

The Nlrp5 genetically-modified mouse strain was obtained from Dr. Lawrence Nelson [43]. Nlrp5 mice were maintained on a mixed 129/C57B16 genetic background and were reared at the Toronto Centre for Phenogenomics (TCP). All animal protocols were approved by the TCP Animal Care Committee and met standards for ethical treatment. Mice were reared in TCP with free access to food and water and were kept on a 12:12 h light: dark cycle.

Nlrp5 breeders: Nlrp5-deficient (homozygous recessive) males were mated with Nlrp5 heterozygous females to produce litters of heterozygous and knockout (homozygous for Nlrp5) pups. Wildtype mice originally derived from heterozygote male and heterozygote female crosses were produced from wildtype females mated to wildtype males. Breeder crosses were ensured to not be between siblings (i.e., used pups born from different litters).

2.2 NLRP5 GENOTYPING Genotypes of Nlrp5 mice were determined by isolating DNA from ear tissue. Protein from ear tissue was first digested overnight at 55⁰C in Tail Lysis buffer, containing 100mM Tris,

5mM EDTA, 200mM NaCl2 and 0.2% SDS, supplemented with Proteinase K (Roche). DNA was extracted using alcohol precipitation, by mixing and pelleting, and was redissolved in 200μl of PCR-grade water. The polymerase chain reaction (PCR) was performed with 2μl of DNA eluent in 20μl total volume of a PCR reaction mix (2μl 10X Taq buffer with (NH4)2SO4, 1.4μl

20mM MgCl2, 0.2μl dNTP, 0.2μl 25μM Nalp5 primer A, 0.2μl 25μM Nalp5 primer B, 0.3μl 25μM Nalp5 primer C, 0.1μl Taq DNA polymerase (Fermentas, Burlington, ON), 13.6μl PCR- grade water). The following primers sequences were used: Nalp5 A [5’-TCA TGT CCT TGG ATG GCA TG-3’], Nalp5 B [5’-ACC GGT GGA TGT GGA ATG TG-3’] and Nalp5 C [5'-

CCA CGT GCT TTC AAG ATT GC-3']. The conditions for each PCR cycle were as follows: denaturation at 95⁰C for 30 seconds, annealing at 59⁰C for 30 seconds, and extension at 72⁰C for 1 minute, with a total of 37 cycles. Amplified products were detected by 1% agarose gel electrophoresis (Bioshop Canada Inc., Burlington, ON, Canada) made with Tris-Acetate-EDTA buffer (Mount Sinai Hospital Media Prep Facility, Toronto, ON) and stained with SYBR Safe DNA gel stain (Invitrogen, Burlington, ON). Bands at 396bp and 242bp corresponded to Nlrp5 wildtype and Nlrp5-deficient genotypes, respectively.

2.3 OOCYTE/ZYGOTE RETRIEVAL

2.3.1 METAPHASE II (MII) OOCYTE COLLECTION Female mice aged six to eight weeks were superovulated by intraperitonial injections of 5IU of Pregnant Mare Serum Gonadotropin (PMSG) (obtained from National Hormone and Peptide Program, NIDDK, Bethesda, MD) to augment the number of mature ovarian follicles, and 46 to 48 hours later with 5IU of human Chorionic Gonandotropin (hCG) (Sigma, Oakville, ON, Canada) to induce ovulation. 14-17 hours after hCG, mice were sacrificed by either cervical dislocation or carbon dioxide gassing, and oviducts were collected in drops of modified Human Tubal Fluid (mHTF, LifeGlobal, Guelph, ON, Canada) media containing HEPES, supplemented with 0.1% Bovine Serum Albumin (BSA) (Sigma). Oocytes were removed from the ampulla of the oviduct with a 26½-gauge needle and were denuded of cumulus cells using hyaluronidase (Sigma) followed by at least 5 washes in mHTF media. Oocytes were either fixed in fixative or were cultured in Human Tubal Fluid media (HTF Xtra, LifeGlobal), supplemented with 0.1% BSA in a humidified incubator at 37⁰C with 5% CO2 until the time of analysis.

2.3.2 GERMINAL VESICLE (GV) OOCYTE COLLECTION Female mice aged six to eight weeks were superovulated with intraperitonial injections of 5IU of PMSG. About 46 to 48 hours after PMSG injection, mice were euthanized by cervical dislocation and ovaries were dissected in drops of mHTF, with 0.1% BSA and 50μM of 3- isobutyl 1-methylxanthine (IBMX) (Sigma) to prevent premature germinal vesicle breakdown. GVs were isolated via the ovary puncture technique, which involves piercing the ovary where follicles are present, avoiding sites of corpus luteum. GV oocytes each posses one distinct nucleus and are surrounded by granulosa cells. Granulosa cells were removed by continuous

34 pipetting up and down using a finely-pulled pipette needle (diameter just greater than diameter of GV oocyte). Denuded oocytes were washed three times in fresh mHTF media supplemented with 0.1% BSA and 50μM IBMX. GVs, in groups of five, were stored in 50μl of guanidinium isothiocyanate (GITC) (5M GITC, 0.5% Sarkosyl, 25mM Sodium Citrate pH 7.0, 20mM 1.4 DTT) and were stored in the -80⁰C freezer for future reverse transcription, real-time PCR studies.

2.3.3 ZYGOTE (FERTILIZED OOCYTE) COLLECTION Female mice aged six to eight weeks were superovulated with 5IU of PMSG by intraperitonial injections, followed 46 to 48 hours later with 5IU hCG injections. After hCG injection, one to two females were placed in a cage containing a single male of proven fertility, overnight, to allow mating. Wildtype females were mated with wildtype males, while Nlrp5- deficient (homozygous) females were mated with Nlrp5-deficient (homozygous) males. About 16 hours after hCG injection, females were inspected for the presence of a vaginal plug, signifying that the female mated. Females were sacrificed by cervical dislocation at 21 (early zygotes) and 26 (late zygotes) hours after hCG injection and oviducts were dissected and placed into drops of mHTF media, supplemented with BSA. Zygotes were flushed out of oviducts using mHTF administered in a blunted 30-gauge needle at the infundibulum. Zygotes were stripped of cumulus cells with hyaluronidase and were washed in multiple drops of mHTF media drops and were fixed.

2.4 WHOLE-MOUNT OOCYTE IMMUNOCYTOCHEMISTRY All incubations were performed in glass-well dishes. Denuded oocytes were washed in mHTF media and were fixed in formalin (Fisher Scientific, Fair Lawn, NJ,USA) or PHEM

(80mM PIPES, 5mM EGTA, 1mM MgCl2, 25mM HEPES at pH of 7.2, 3.7% Formaldehyde, 10% Triton X-100) for 1 hour. PHEM fixative contains a detergent which bleaches out non- permanently bound structures and has lower cytoplasmic staining. Oocytes were then washed in Phosphate Buffered Saline (PBS) for 5 minutes, followed by a 40 minute incubation in Nuclear permeabilization solution (0.4% Surfact-Amps X-100 (Pierce, Nepean, ON) in Antibody Diluent (0.004% Sodium Azide and 0.0001% Gelatin in PBS) at room temperature, followed by a 1 hour incubation in Blocking solution (0.5% BSA in Antibody Diluent). Primary antibody was diluted in Primary Antibody Diluent (0.1% BSA in Antibody Diluent) and oocytes were

35 incubated in 10μl of diluted primary antibody in a Terasaki plate (Nunc Brand, Roskilde, Denmark) overnight at 4˚C, in a wet chamber, to prevent evaporation. After primary antibody incubation, oocytes were washed for 10 minutes in Washing Solution (0.1% Triton-X in PBS), three times. Following which, oocytes were incubated in secondary antibody diluted in Secondary Antibody Diluent (0.1% Triton-X and 0.0001% Gelatin), in the dark, for one hour. Oocytes were washed in PBS for 10 minutes, three times, in the dark and were transferred onto a slide and excess PBS was removed. Slide was placed in coplin jar containing 1μg/ml of (4,6- diamidino-2-phenylindole) DAPI (Sigma) for 10 minutes. Oocytes were mounted with 50% glycerol (in PBS) and a coverslip was added to protect sample and sealed with nail polish. Slides were stored at 4˚C until time of analysis under the microscope.

Table 4: List of Primary Antibodies and Dilutions

Primary Antibody Source Catalogue # ICCH Fixative Dilution Anti-NALP5 Produced by Dr. Jurrien Dean - 1:15000 PHEM Published in: [47] Elute BRCA1 1904 0.13μg/μl Produced by Dr.Razqallah Hakem - 1:600 PHEM H79 (p73) polyclonal-rabbit Ab Santa Cruz sc-7957 1:100 PHEM Santa Cruz Biotechnology, Santa Cruz, CA, U.S.A. α-tubulin (bovine) mouse Molecular Probes, Invitrogen 236-10501 1:2000 PHEM monoclonal IgG anti-Bub1b, N-Terminal (Ab-3). Calbiochem PC619 1:200 PHEM Mouse (Sheep) Dynactin p50 mouse IgG1 BD Transduction Laboratories 611002 1:50 PHEM Aurora 2. AK rabbit Cell Signaling 3092 1:100 PHEM Anti-murine BUB1 monoclonal Produced by Dr. Frank McKeon 4B12 1:200 PHEM Published in: [88] purified mouse IgG1 Anti-AIM- BD Transduction Laboratories 611082 1:300 PHEM 1 Pericentrin rabbit Covance PRB-432C, 1:500 PHEM LN#142037002 polyclonal rabbit Survivin Novus Biologicals NB500-201 1:500 Formalin polyclonal rabbit 53BP1 Novus Biologicals NB100-904 1:1000 Formalin Phospho-Histone H2A.X Cell Signaling 9718 1:500 Formalin (Ser139) Rabbit mAb Rabbit phospho-Histone H3 Cell Signaling 9701S 1:300 Formalin (Ser10) α-2b tri meth H3(aa5-15) K9; 3.1 Produced by Dr. Thomas Jenuwein - 1:400 Formalin mg/mL Published in: [129] [lysine-9 tri-methylated histone H3] BUB1 (H-300) rabbit polyclonal Santa Cruz sc-28257 1:400 PHEM BUBR1 (C-20) goat polyclonal Santa Cruz sc-16195 1:500 PHEM Rabbit-Mouse Aurora C – Bethyl Laboratories A400-023A 1:200 PHEM BL1217 anti-ATM protein kinase Rockland 200-301-400 1:300 Formalin pS1981, mouse monoclonal

Table 5: List of Secondary Antibodies and Dilutions

Secondary Antibody Source Catalogue ICCH # Dilution Alexa Fluor® 488 rabbit anti-mouse Molecular Probes, A11059 1:600 IgG (H+L) 2 mg/mL Invitrogen Alexa Fluor® 594 donkey anti-rabbit Molecular Probes, A21207 1:600 IgG (H+L) 2 mg/mL Invitrogen Alexa Fluor® 488 goat anti-mouse Molecular Probes, A11001 1:600 IgG (H+L) 2 mg/mL Invitrogen Alexa Fluor® 488 donkey anti-goat Molecular Probes, A11055 1:600 IgG (H+L) 2 mg/mL Invitrogen

2.4.1 NEGATIVE CONTROLS AND BLOCKING PEPTIDE Negative controls were oocytes/zygotes that were not exposed to primary antibody, but were stained with secondary antibody. All secondary antibodies have been tested as negative controls and were titrated to have no detectable signal in metaphase II-arrested oocytes, to ensure that there was no cross-reactivity of the secondary antibody in the sample.

Aurora C antibody recognizing the BL1217 epitope, mapping the region between residue 250 and the C-terminus (residue 282) was used with a blocking peptide. In 200μl of antibody diluent, 1μl of Aurora C antibody was added, along with 5μl of the peptide. Mixture was incubated overnight at room temperature. Prior to incubating oocytes in primary antibody, the peptide mixture was centrifuged for 15 minutes and was then incubated overnight at 4˚C in a wet chamber, following the same immunocytochemistry protocol.

Optimal controls for many of the evaluated proteins are through analysis in mouse models that have the gene knocked-out. It has previously been shown that the TAp73 protein decreases in TAp73-deficient oocytes [91]. Likewise, I also analysed NALP5 expression in Nlrp5-deficient oocytes [Figure 13A].

2.5 NOCODAZOLE TREATMENT Nocodazole is a pharmacological agent and spindle toxin commonly used to induce spindle damage, by means of interfering with microtubule polymerization. Nocodazole induces metaphase arrest, since it causes depolymerisation of spindles, activating the SAC, ultimately causing the arrest of the cell cycle at metaphase of mitosis or meiosis [103]. Nocodazole dose

38 was determined by testing out various concentrations of nocodazole that would produce the effect of a disrupted spindle. Tested concentrations include: 5μg/ml, 10μg/ml and 20μg/ml of nocodazole. 10μg/ml of nocodazole was determined to be the optimized, lowest dose that triggered the depolymerised-spindle phenotype.

10μg/ml of nocodazole (originally diluted in DMSO), was diluted pre-equilibrated HTF medium and was incubated for 15 minutes. Vehicle control oocytes were incubated for 15 minutes in the same concentration of DMSO 10μl in 1ml of HTF media. After treatment, oocytes were washed in HTF medium and were fixed in PHEM or formalin fixative for immunofluorescence staining.

2.6 UV IRRADIATION AND OOCYTES CULTURE Oocytes stripped of cumulus cells were added to a 35mm dish containing pre-equilibrated HTF medium. Stratalinker UV Crosslinker 1800 was used to induce ultraviolet radiation at a dose of 120,000 microjoules/cm2 or [12 J/m2]. UV was delivered using 254-nm UV light bulbs (8 watts each), which is also known as Ultraviolet type C (UVC) or UV short wave. Both the UV-treated and non-treated groups were placed in the Cosslinker, however, the non-exposed, control group was shielded from radiation (i.e., was covered with a lid), while the UV-treated group was uncovered. After UV exposure, UV-exposed and non-exposed oocytes were cultured in HTF medium in the incubator at 37⁰C with 5% CO2 for one and five hours to see effects of DNA repair. The dose of UV irradiation was based on previous work in somatic cells, as the dose that induced G2/M phase arrest [130]. Timepoints were chosen from literature searches on somatic cells [122, 131] and this was modified to 5 hours because oocytes can undergo post- ovulatory aging if left in culture for a prolonged period of time after ovulation, creating other confounding factors. Following culturing, control and treated oocytes were fixed in PHEM or Formalin and immunocytochemistry for various epitopes was performed.

2.7 COMET ASSAY The Comet Assay (Trevigen 4250-050-K, Gaithersburg, MD, USA) is an experiment that measures the amount of DNA damage that a cell endures. We wanted to assess whether Nlrp5- deficient oocytes exhibited altered levels of DNA damage. Ovulated oocytes collected 14-17 hours after hCG injection were retrieved by piercing the ampulla of the oviducts and oocytes

39 were stripped off of cumulus cells and were used for this assay. MII oocytes were ensured to have no cumulus cells since cumulus cells would produce DNA contamination. Stripped- oocytes were added to a drop of melted Low Molten (LM) agar (cooled to 37⁰C) on the Comet slide, such that oocytes were arranged perpendicularly relative to the direction of electric field so that comet tails would not conjoin. Additional LM agar was added to seal the gel drop onto the slide. Lysis buffer was added to the slide on ice in dark for 35 minutes, followed by alkaline unwinding solution (200mM NaOH, 1mM EDTA) incubation for 30 minutes. Slides were then washed in TBE buffer for five minutes, two times. Slides were run for five minutes at 30V and 5-9mA in TBE buffer. Slides were fixed in ice cold methanol, followed by ice cold ethanol. Slides were dried and stored at room temperature until analysis. Slides were stained with a dilution of 1:10,000 of SYBR Safe (diluted in TBE). Samples were imaged and the length of the comet tail was visualized using Volocity software. A longer length of the comet tail signifies increased DNA damage compared to no tail [132].

2.8 IMAGING: FLUORESCENCE MICROSCOPY Zygotes, misaligned chromosomes, and abnormal spindle samples were serially scanned on the deconvolution microscope (Olympus IX70, Applied Precision Inc., Issaquah, WA, USA) and images were analysed using DeltaVision software (Applied Precision Inc, Issaquah, WA, USA). Oocytes and zygotes were imaged using the 20X objective under either: fluorescein isothiocyanate (FITC), rhodamine isothiocyanate (RITC) and DAPI (4', 6-diamidino-2- phenylindole) filters.

The immunostained cells were serially scanned using the Quorum WaveFX Spinning Disk Confocal Laser-Scanning Microscope system (Leica DMI6000B; Quorum, Guelph, ON). Oocytes were imaged using the 40X (dry) or 100X (oil) objective with the Green Fluorescent Protein, Red Fluorescent Protein and DAPI channels. Depending on localization of the protein, various regions of the oocyte were imaged, such as the global oocyte, the spindle, or the chromosomal plate. 40X images were optically sectioned with a spacing of 0.4μm and a total z- distance of 9μm, while 100X images had a spacing of 0.2μm and a total z-distance of 7μm. Both wildtype and Nlrp5 knockout samples were imaged at the same time to reduce the effects of fluorescence decay. Exposure times were kept consistent between treatment groups per experiment. The exposure time was calculated by sampling about five cell samples

(oocytes/zygotes) per treatment group (i.e., wildtype and knockout) and calculating the average exposure time. Then the lowest average exposure time was used to acquire images of all samples from all of the treatment groups. The secondary antibody alone (negative) control was used qualitatively to ensure that there was no non-specific binding of the secondary antibody to the sample.

Volocity version 5.2.2 (Improvision Ltd., Coventry, UK) software was used for quantification. For global quantification of fluorescence for the entire MII oocyte, the percent intensity protocol was used targeting the region of the entire oocyte. For quantification of florescence levels on chromosomal plate, regions of interest were selected using DAPI as the target region. Spindle poles localization of BRCA1 was quantified using the percent intensity protocol, selecting the spindle pole region. For all quantification measures, the mean fluorescence intensity values were used.

2.9 MICROARRAY Microarray analysis was performed on 100 metaphase II-arrested oocytes per sample from Nlrp5-deficient and wildtype females, aged six to eight weeks. Four sets of samples were collected per genotype. Samples were collected in Trizol (Invitrogen, USA) and RNA extraction was performed using Trizol with glycogen as a carrier. Two rounds of amplification were performed prior to hybridization. Used Affymetrix chips (Mouse MOE 430 2.0) covering the full genome. Investigated targets with fold change greater than 1.76 that passed the t-test with p≤0.05.

Gene profiling analysis was done to determine the functional links among the identified target genes, using the Significant Analysis of Mean (SAM) algorithm and genes that had a 2- fold difference were mapped to I2D (protein-protein interaction) database to see what pathways are affected. This was done in collaboration with Dr. Jurisica.

2.10 STATISTICAL ANALYSIS All data, comparing wildtype (WT) and Nlrp5-deficient (knockout (KO)) oocytes, was transformed relative to wildtype fluorescence intensity levels. This was done to allow for the compilation of datasets from various days to increase the sample size. For the transformed data,

41 the mean fluorescence intensity for wildtypes was expressed equivalent to one and that belonging to Nlrp5-deficient samples was expressed in relation to wildtypes plus/minus standard error (SEM), and the number of oocytes that were quantified for fluorescence level was labelled in parentheses as (n= ). For each experiment, oocytes/zygotes were collected and pooled from various mothers to ensure that there was a representative sample population with variation and to ensure that the potential observed effect would not be due to that of a single mother. For all of the analysis of spindle checkpoint protein expression and UV damage marker expression, we only used normal appearing metaphase II (MII) oocytes for quantification. Particularly for spindle morphology evaluation, we only selected the samples that appeared to have normally aligned chromosomes and spindles that were suitably oriented (i.e., longitudinal orientation). For instance, if the spindle was malformed the proteins were not considered for quantitative purposes, they were only used for qualitative analysis. This was done because if a difference was observed in protein expression between treatment groups (i.e., WT and Nlrp5 KO or between UV-exposed and non-UV-exposed) of normal-looking oocytes, then it could be inferred that the effect of the proteins or chromosomes displaying an abnormal phenotype could be a consequence of the difference in the protein expression. Following the selection of samples, we used the mean fluorescence intensity value to account for the size in voxels of the targeted region. This was done to ensure that the values that we scored were not inaccurately skewed.

Frequency of abnormal pronuclear number in zygotes, frequency of abnormal spindle morphology in oocytes and frequency of chromosomal misalignment/breaks were tested for significance using the Chi-squared (χ2) test. Results were considered statistically significant when p<0.05.

All UV baseline values (UV-exposed and non-exposed and 1h and 5h timepoints) were analysed by Two-way Analysis of Variance (ANOVA), using the Holm-Sidak method for pairwise multiple comparisons. p<0.05 was considered statistically significant.

All other data: If data had a) a normal distribution and b) equal variance, then data were analysed using unpaired samples, two-tailed (95% Confidence Interval) t-tests to compare against two groups (i.e., WT and Nlrp5 KO). If data did not have either a normal distribution or had unequal variance, then a non-parametric Mann-Whitney Rank Sum test was performed. p<0.05 was considered statistically significant. All statistical analysis was conducted using SigmaPlot Version 11, Systat Software Inc. (San Jose, CA, USA). Graphs were created using Prism 4, GraphPad Software Inc. (La Jolla, CA, USA).

3 RESULTS

3.1 HISTONE METHYLATION PATTERN IN NLRP5-DEFICIENT ZYGOTES Histone modifications and chromatin remodelling complexes are required for proper zygotic genome activation during preimplantation embryo development [133]. Tri-methylation of lysine 9 of Histone 3 (H3K9me3) contains the dominant maternal transgenerational signal for pericentric heterochromatin formation [133]. To determine if there was a difference in histone methylation patterns in Nlrp5-deficient mice relative to wildtype mice, an antibody recognizing H3K9me3 was used on zygotes and was analyzed using deconvolution immunofluorescence microscopy [Figure 9]. In wildtype zygotes, only the maternal genome is H3K9me3 methylated and H3K9me3 is always absent from male chromatin [134], causing H3K9me3 to be a good marker of maternal DNA. We analysed wildtype zygotes (produced from wildtype mothers mated with wildtype fathers) and Nlrp5-deficient zygotes (bred from Nlrp5 KO mothers and Nlrp5 KO fathers) at two time points of development, early (21 hours post hCG injection) and late (26 hours post hCG injection) zygotes. The maternal pronuclei of both knockout (n=30) and wildtype (n=29) zygotes were methylated, while the paternal pronuclei remained unmethylated. Similarly, in late zygotes, in both wildtype (n=20) and in knockout (n=28) zygotes, the maternal pronuclei were methylated and the paternal pronuclei were unmethylated. While investigating methylation patterns or intensity in Nlrp5-deficient and wildtype mouse zygotes, no significant differences were observed at both time points.

Early Zygotes Late Zygotes WT Nlrp5 KO WT Nlrp5 KO PB DNA PB PB ♀ ♀ ♂ ♀ ♀ ♂ ♂ ♂ PB

H3K9me3

n=29 n=30 n=20 n=28 Figure 9: Methylation pattern in Early and Late Nlrp5-deficient zygotes.

Trimethylation of Lysine 9 on histone H3 (H3K9me3) methylation pattern in zygotes (1-cell embryos). Indirect immunocytochemistry (ICCH) was performed on early zygotes (collected 21 hours post hCG injection) and late zygotes (collected 26 hours post hCG injection) using H3K9me3 antibody. Images were acquired using a deconvolution microscope. The methylation profile does not seem to be altered between Nlrp5 knockout (KO) and wildtype (WT) zygotes at both early and late zygote stages. DAPI-green; H3K9me3 (rabbit)-red. PB-Polar Body.

3.2 THE ROLE OF NLRP5 IN THE ACTIVATION OF THE SPINDLE ASSEMBLY

CHECKPOINT IN METAPHASE II OOCYTES

3.2.1 PRONUCLEAR NUMBER ABNORMALITIES IN NLRP5-DEFICIENT ZYGOTES While examining the nuclear staining of zygotes with DAPI by deconvolution fluorescence microscopy, an interesting observation was made. A normal zygote consists of two parental pronuclei: one composed of the maternal DNA from the oocyte and the other composed of paternal DNA from the sperm. During our assessment of histone methylation pattern we observed a greater frequency of abnormal pronuclear number in both early and late Nlrp5- deficient zygotes. The pronuclear number abnormalities included: three pronuclei of comparable size (observed in 40% of KOs), two pronuclei plus a single micronucleus (observed in 40% of KOs), or zygotes with a single pronucleus (observed in 20% of KOs) [Figure 10A]. There was an 8% occurrence of abnormal pronuclei number in wildtypes (n=51) and a significantly higher frequency in Nlrp5-deficient zygotes (35%; n=57) ( 2= 11.56, p<0.001) [Figure 10B].

Since H3K9me3 is a marker of the maternal genome, we analysed the origin of multinucleated DNA using the H3K9me3 antibody. We observed one unmethylated, paternal pronucleus while the remaining pronuclei were H3K9me3 positive, indicating their maternal origin [Figure 10C]. This suggests that the extra pronucleus or micronucleus is a result of defective meiosis that originated in the oocyte. Furthermore, previous observations in our laboratory looked at multinucleation in 4 to 8-cell stage embryos and found that about 30% (n=13/39) of Nlrp5 knockouts had blastomeres with multiple nuclei compared to only 4% (n=1/27) of wildtype embryos (Jurisicova and Fernandes, 2011). Because of this increased multinucleation phenotype in Nlrp5-deficient zygotes and preimplantation embryos, which likely resulted from abnormal meiosis, we focused the following experiments on the analysis of meiotic defects in ovulated oocytes.

A 3PN Micronucleus 1PN

DAPI

PB PB

B C 2=11.56, p<0.001 40 DAPI ♂ PB 30 ♀

♀ ♂ 10 H3K9me3

Zygotes with Zygotes Abnormal n=51 n=57 PB ♀ Number Number of Pronuclei (%) 0 Wildtype (WT) Nlrp5 Knockout (KO)

Figure 10: Pronuclear Abnormalities in Nlrp5-deficient zygotes.

A: Examples of pronuclear number abnormalities in zygotes (1-cell embryos). Zygotes were stained with DAPI (blue), highlighting nuclear DNA and were imaged using a deconvolution microscope. Pronuclear abnormalities include: presence of 3 pronuclei (3PN), presence of a micronucleus, and presence of a single 1 pronucleus (1PN). PB-Polar Body. B: A significantly increased number of Nlrp5-deficient zygotes displayed abnormalities in pronuclei numbers compared to wildtype zygotes (p<0.001). Chi-squared (χ2) test. Values are expressed as percent frequency. (WT-Wildtype, KO-Knockout) C: Immunocytochemistry with H3K9me3 Methylation antibody, marker of maternal DNA, showing that extra pronucleus was maternal in origin. H3K9me3 (rabbit)-red; DAPI-green. PB- Polar Body.

3.2.2 SPINDLE MORPHOLOGY AND CHROMOSOMAL MISALIGNMENT ABNORMALITIES IN

OVULATED, METAPHASE II OOCYTES As multinucleation has been linked to improper cell division and chromosome missegregation, this directed us to investigate the cellular structure that is responsible for accurate chromosomal segregation – the spindle and its morphology in ovulated oocytes. Immunocytochemistry was performed using an α-tubulin antibody, recognizing spindle microtubules, counterstained with DAPI (DNA) [Figure 11]. Spindle morphology abnormalities that were observed, include: wide spindles, two connected spindles, spindles that exhibited unequal tension [Figure 11.A.b], spindles with microtubules not attached to the kinetochore of chromosomes [Figure 11.A.c], mono-polar spindles, multi-polar (more than 2 poles) spindles [Figure 11.A.d]. While scoring spindle morphology between spindles of wildtype and Nlrp5- deficient MII-arrested oocytes, we noticed that about 5% of wildtype (n=79) oocyte spindles have abnormalities, compared to about 28% (n=67) of abnormalities seen in Nlrp5-deficient spindles. The frequency of spindle morphology abnormalities was statistically significant between wildtype and knockout oocytes ( 2=14.82, p=0.0001).

Negative Control A

α-Tubulin DAPI PB a a b e

b c d B 2=14.82, p=0.0001 30

10 Morphology (%)Morphology

Percentage of Ovulated Percentage n=79 n=67 0 WT Nlrp5 KO Oocytes withOocytes Abnormal Spindle Figure 11: Proportion of ovulated oocytes with abnormal spindle morphology.

A: Examples of abnormal spindle morphology. Immunocytochemistry was performed on MII (metaphase of meiosis II) oocytes and images were captured using laser-scanning confocal microscopy. α-Tubulin (mouse-green) stains meiotic spindles and DNA is stained with DAPI (blue). a: show a normal, barrel-shaped spindle, whereas b-d: are spindles of poor morphology. b: a spindle that exhibited unequal tension, evidenced by lagging chromosome (arrowhead), c: detached spindle (arrow), d: multi-polar spindle (arrows) e: Negative control MII oocyte is exposed to secondary antibody only, goat anti-mouse green and DAPI. B: Percentage of abnormal spindles in Nlrp5 wildtype and Nlrp5-deficient MII oocytes. Significant increase in spindle morphology abnormalities were observed in Nlrp5-deficient oocytes compared to wildtypes (p=0.0001). Values are expressed in percentage. Chi-squared χ2 test.

The increased prevalence of multinucleation in zygotes and abnormal spindles in Nlrp5- deficient oocytes led us to investigate the chromosomal status. There was a large degree of variability of chromosomal misalignment, ranging from subtle cases, of single chromosomes lagging or potentially breaking, to severe cases of chromosomal scattering [Figure 12]. We found that in about 3% of wildtype oocytes (n=125), chromosomes were improperly aligned, whereas this was observed in about 22% of knockout oocytes (n=125). When compared between wildtype and Nlrp5-deficient oocytes, there was a significant increase in knockouts displaying a greater frequency of misaligned chromosomes ( 2=20.64, p<0.0001). These data imply that Nlrp5 is required for proper meiotic progression and chromosomal segregation, as well as maintaining genome integrity.

DAPI

a b c

B 2  =20.64, p<0.0001 25

5 Percentage of Ovulated Percentage Oocytes withOocytes Abnormal n=125 n=125 0 Chromosomal Chromosomal Alignment (%) WT Nlrp5 KO

Figure 12: Proportion of ovulated oocytes with misaligned chromosomes.

A: Examples of Chromosomal Alignment Abnormalities. Immunocytochemistry was performed on MII oocytes and images were captured using laser-scanning confocal microscopy. DNA is stained with DAPI (blue). a: chromosomes aligned at the metaphase plate, b-c: are MII chromosomes with varying degrees of misalignment. Arrows denote lagging or misaligned chromosomes. B: Percentage of abnormal chromosomal alignment in Nlrp5 wildtype and knockout MII oocytes. A significant higher frequency of chromosomal misalignment was observed in Nlrp5- deficient oocytes compared to wildtypes (p<0.0001). Values are expressed in percentage. Chi- squared χ2 test.

3.2.3 NALP5 LOCALIZATION IN MII OOCYTES To investigate the localization of NALP5 in metaphase-II oocytes, we performed immunocytochemistry using an anti-NALP5 antibody and anti-α-tubulin. The NALP5 antibody was used to stain oocytes in a previous publication [47]; all publications describing MATER localization in ovulated oocytes have only described cytoplasmic and subcortical staining [47, 64]. We used a different fixative used from past publications. PHEM fixative, composed of a cytoskeletal stabilization buffer, retains components bound to the cytoskeleton and soluble proteins are washed away. We observed that NALP5 appeared to localize cytoplasmically with enrichment on the chromosomal DNA region in 71% of oocytes (n=28) [Figure 13A, B]. In Nlrp5-deficient oocytes, NALP5 expression was absent or greatly reduced on the chromosomal plate (n=43), indicating specificity of the antibody. When these oocytes were co-stained with kinetochore marker, dynactin p50, there was some degree of colocalization [Figure 13C]. Interestingly, dynactin p50 and NALP5 were only visible on the outer edge of the chromosomal plate, with hardly visible staining in the core of the chromosomal region. This may have been caused because the kinetochores may have been hidden or were inaccessible by the antibodies. One future experiment could be to try chromosomal spreads to allow the kinetochores to be completely visible and accessible to antibodies.

A WT Nlrp5 KO Negative Control a b c α-Tubulin NALP5 DAPI

NALP5

40x

Dynactin p50 B WT C DAPI NALP5 NALP5 Dynactin p50

NALP5 WT DAPI

100x

NALP5

100x Figure 13: Localization of NALP5 protein in MII oocytes.

A: Using laser-scanning confocal microscopy. Oocytes were fixed in PHEM fixative and indirect immunocytochemistry was performed using an NALP5 antibody. In wildtype oocytes a: NALP5 localizes cytoplasmically with enrichment on the chromosomes (circled-region), whereas b: in Nlrp5-deficient oocytes there is drastically reduced chromosomal staining. NALP5 (rabbit)-red; α-Tubulin (mouse)-green; DAPI-blue. PB-Polar Body. c: Negative control is exposed to secondary antibody only. Top image – merged, goat anti-mouse green and donkey anti-rabbit red; bottom image – anti-rabbit red; DAPI-cyan.

B: 100X magnification on the chromosomal plate of wildtype MII oocytes. NALP5-red; DAPI- cyan.

C: Immunocytochemistry in wildtype MII oocytes with NALP5 and Dynactin p50, kinetochore marker, shows that NALP5 and Dynactin p50 coincide on the kinetochores. 100X magnification on the chromosomal plate. NALP5 (rabbit)-red; Dynactin p50 (mouse)-green; DAPI-blue.

3.2.4 SPINDLE ASSEMBLY CHECKPOINT ANALYSIS IN NLRP5 DEFICIENCY AT THE MII

OOCYTE-STAGE Due to the observed phenotypes of multinucleation and spindle defects, implicated with abnormal chromosomal segregation, we wanted to assess the relationship between Nlrp5 and the Spindle Assembly Checkpoint. To do this, we performed immunofluorescence microscopy on ovulated oocytes and analysed samples using the spinning disc confocal microscope. SAC- involved proteins that we investigated include: BUB1, BUBR1, TAp73, Aurora kinase B, Aurora kinase C and Dynactin p50.

3.2.4.1 BUB1 PROTEIN EXPRESSION Protein kinase BUB1 (Budding Uninhibited by Benzimidazole) is a SAC protein that localizes to the kinetochores and is necessary for the proper attachment of kinetochores to microtubules [83]. It was recently reported that Bub1 knockouts are embryonic lethal and that Bub1 heterozygous mouse oocytes were aneuploid (76%), due to premature sister chromatid separation [87]. In our immunocytochemistry experiment, staining on the kinetochores was visualized as punctate foci associated with the chromosomes. While this pattern was observed in about 83% of wildtypes (n=23), only 24% of Nlrp5 knockouts (n=34) showed this type of staining [Figure 14]. However, it should be noted that the knockouts that did show the presence of punctate staining on the kinetochores were very faintly stained compared to the pattern observed in wildtypes. Quantification of BUB1 fluorescence intensity on the metaphase plate [Figure 5A] of the spindle (the area where the chromosomes align during metaphase II) revealed that BUB1 staining was decreased in Nlrp5-deficient oocytes (0.81 ± 0.047, n=34) relative to wildtype (1.00 ± 0.037, n=20) oocytes (t test, p=0.005). This data suggests that due to Nlrp5 deficiency, BUB1 is failing to be recruited to the kinetochores and consequently, Nlrp5 affects BUB1’s role in maintaining the kinetochore-bound scaffold. This pattern of expression was verified using two different BUB1 antibodies. The BUB1 negative controls for this experiment, followed the same immunocytochemistry protocol, but were not exposed to BUB1 antibody and did not show any localization on the kinetochores, suggesting that the secondary antibody did not contribute to the pattern of observed staining.

BUB1 WT DAPI Nlrp5 KO

BUB1 ** 1.1 (p = 0.005) 1.0 0.9 0.8 0.7 0.6 0.5 (RFU) 0.4 0.3 0.2 Chromosomal Chromosomal Plate 0.1 Relative Expression on Relative Expression n=20 n=34 0.0 WT Nlrp5 KO

Figure 14: BUB1 Protein expression in MII oocytes.

Immunocytochemistry was performed in MII oocytes to investigate the protein level of BUB1. Confocal microscopy images on the chromosomal plate. BUB1 Antibody (rabbit)-red; DAPI- blue. In wildtype (WT) oocytes, BUB1 localizes to the kinetochores and in Nlrp5-deficient oocytes this staining is absent. Quantification of fluorescence intensity suggests that there is a significant decrease in BUB1 protein levels on the metaphase plate of Nlrp5-deficient oocytes compared to wildtype oocytes (p=0.005). Values represent fluorescence intensity relative to WT levels ± SEM. Asterisks denote means are significant. Mann-Whitney Rank Sum test.

3.2.4.2 BUBR1 PROTEIN EXPRESSION BUBR1 is a SAC protein with a defined role in mitosis and has recently been identified as a SAC protein involved in meiosis, more specifically oogenesis [88]. Decrease in BubR1 (hypomorphic allele) causes oocytes to have abnormal MII configuration and causes mouse embryonic fibroblasts to undergo premature sister chromatid separation [90]. Using a BUBR1 antibody, immunocytochemistry was conducted on MII oocytes. Expression analysis revealed that wildtype oocytes (n=31) showed localization of BUBR1 to kinetochores, while Nlrp5- deficient MII oocytes showed decreased staining around the DNA (n=30) [Figure 15]. When the fluorescence intensity of BUBR1 was quantified, there was a significant decrease in BUBR1 expression in Nlrp5-deficient (0.78 ± 0.05, n=30) oocytes compared to wildtypes (1.00 ± 0.06, n=31) (p=0.006). This expression and localization of BUBR1 was verified using two different antibodies, from different sources. Hence, taken together, Nlrp5 deficiency compromised the localization of two SAC proteins, BUB1 and BUBR1, at the kinetochores. Moreover, this finding is consistent with the BUB1 protein expression pattern, in that the lack of a proper kinetochore scaffold does not allow for BUBR1 attachment to kinetochores as well.

BUBR1 2⁰ Ab Only WT DAPI Nlrp5 KO Control

BUBR1 1.1 ** 1.0 (p = 0.006) 0.9 0.8 0.7 0.6 0.5 (RFU) 0.4 0.3 0.2 Chromosomal Chromosomal Plate 0.1 Relative Expression on Relative Expression n=31 n=30 0.0 WT Nlrp5 KO

Figure 15: BUBR1 Protein expression in MII oocytes.

Immunocytochemistry was performed and confocal microscopy was used to investigate the protein level of BUBR1. Images were focused on the chromosomal plate of MII oocytes. BUBR1 Antibody (goat)-green; DAPI-cyan. In wildtype oocytes, BUBR1 localizes to the kinetochores and in Nlrp5-deficient oocytes this staining is absent. Negative control is exposed only to donkey anti-goat secondary antibody. Quantification of fluorescence intensity on chromosomal plate suggests that there is a significant decrease in BUBR1 protein levels in Nlrp5-deficient oocytes compared to wildtype oocytes (p=0.006). Values represent fluorescence intensity relative to wildtype levels ± SEM. Mann-Whitney Rank Sum test. Asterisks denote means are significant.

3.2.4.3 P73 AND TAP73 PROTEIN EXPRESSION Since we previously saw some phenotypic similarities in TAp73-null and Nlrp5-null oocytes and embryos, such as spindle abnormalities in MII oocytes and greatly reduced percentage of embryos that reached the 4-8 cell, morula, and blastocyst stages [44], we decided to investigate whether there was interplay between the two proteins. Previous data from our laboratory indicates that total p73 transcript is statistically significantly decreased in Nlrp5- deficient (0.37 ± 0.06, n=9) compared to wildtype (1.04 ± 0.10, n=10) MII oocytes (p<0.0001, t- test) (Perumalsamy et al. 2011, unpublished). These findings led us to carry out immunostaining of p73 protein in oocytes, using the H79 antibody. Quantification of global p73-staining revealed that there was a significant decrease in fluorescence intensity in Nlrp5-deficient oocytes (0.83 ± 0.05, n=7), compared to wildtype (1.00 ± 0.02, n=14) oocytes (p=0.004, Mann- Whitney test) [Figure 16A]. With further analysis using the same H79 antibody on the chromosomal plate, we found that in wildtype MII oocytes, we see higher background cytoplasmic staining on the chromosomal plate in wildtypes [Figure 16B]. We also found that in wildtype oocytes, p73 localized to the spindle and chromosomes, which is consistent with the localization pattern of the TA-specific isoform of p73, TAp73 [91]. More specifically, there is enrichment on the spindle region closer to the chromosomes as well as on the kinetochores. Nlrp5-deficient oocytes have reduced cytoplasmic staining and p73 fails to localise to the spindle region in a majority of oocytes. In wildtype oocytes, 9/13 oocytes showed expression of p73 on the spindles. While all Nlrp5-deficient oocytes (18/18) had expression of p73 on the chromosomes, only 3/18 had staining on the spindle. This data indicates that TAp73 protein localization to the spindle is altered in Nlrp5-deficient MII oocytes. These observations are consistent with the findings of Tomasini and others’ which showed that TAp73 regulates the activity of the SAC by modulating the activity and localization of BUBR1 [91]. Thus, with Nlrp5 deficiency we see perturbed levels of TAp73, BUBR1 and BUB1, causing altered kinetochore scaffold and function of the SAC.

A p73 1.1 ** 1.0 (p = 0.004) 0.9 0.8 0.7 0.6 0.5 0.4 0.3

0.2 Global Oocyte Global (RFU) Oocyte

Relative Expression in Relative Expression 0.1 n=14 n=7 0.0 WT Nlrp5 KO WT Nlrp5 KO B p73 DAPI

p73

69% Spindle with chromosomal localization 17% Spindle with chromosomal localization of p73 (9/13) of p73 (3/18)

Figure 16: p73 and TAp73 expression pattern in MII oocytes.

A: Immunocytochemistry was performed in MII oocytes to investigate the protein level of total p73. Quantification of fluorescence (measured in Relative Fluorescence Units) intensity of the global oocyte, indicative of total p73 levels, suggests that there is a significant decrease in total p73 protein levels in Nlrp5-deficient oocytes compared to wildtype MII oocytes (p=0.004). Values represent mean fluorescence units relative to wildtype expression ± SEM. Mann- Whitney Rank Sum test.

B: TAp73 protein levels were investigated by the staining of p73 that is specific to the spindles. In wildtype MII oocytes there is evident spindle staining (boxed region), however in Nlrp5- deficient oocytes staining on the spindle was absent, but was only present on the chromosomes, suggesting that there is decreased TAp73 protein in Nlrp5-deficient oocytes. p73 (H79) Antibody (rabbit)-red, DAPI-cyan.

3.2.4.4 AURORA KINASE B Aurora kinases are involved in chromosomal segregation and regulate chromatin status via phosphorylation of histones [Figure 17A].

Aurora B is essential for proper chromosome segregation, kinetochore-microtubule interaction and cytokinesis [94]. It has previously been shown that by microinjecting an AuroraB-tagged GFP protein into germinal vesicle oocytes and allowing for in vitro maturation, they found that Aurora B was enriched on the centromeres or kinetochores [92]. We analysed Aurora B protein expression in MII oocytes by immunocytochemistry and found that, consistent with previous reports [92, 97], wildtype oocytes had low to undetectable levels of endogenous Aurora B [Figure 17B]. Interestingly, Nlrp5-deficient oocytes (1.20 ± 0.07, n=18) had much higher levels of Aurora B on kinetochores than wildtypes (1.00 ± 0.05, n=15). This suggests that Aurora kinase B protein levels are increased in Nlrp5 deficiency.

3.2.4.5 AURORA KINASE C We next decided to look at the expression of Aurora C kinase (germ cell-specific Aurora kinase), since there was differential expression between wildtype and Nlrp5-deficient oocytes in our microarray analysis [Table 6]. Moreover, it has recently been shown that in mouse oocytes, Aurora C deficiency causes cytokinesis failure in meiosis I and the production of large polyploid oocytes [97]. It has also been reported that Aurora C localizes to the centromeres in metaphase II oocytes. By performing immunocytochemistry and quantifying the fluorescence intensity of Aurora C, we found that Aurora C localized to the centromeres in wildtype oocytes (1.00 ± 0.05, n=16) [Figure 17C]. Similar to Aurora B kinase, Nlrp5-deficient oocytes displayed significantly increased expression of Aurora C at the centromeres (1.43 ± 0.08, n=20) (p=0.0001). A blocking peptide was used as a negative control and showed no staining on the centromeres, confirming specific reactivity of this antibody.

3.2.4.6 DYNACTIN P50 PROTEIN EXPRESSION The final SAC protein that we assessed the protein expression of was the cytoskeletal, motor protein dynactin p50. In metaphase of mitosis in somatic cells, it has been shown that dynactin p50 disappears from the kinetochores upon its alignment at the metaphase plate, but is very bright in kinetochores of nonaligned metaphase chromosomes [101]. Our immunofluorescence microscopy analysis revealed that dynactin p50 localized to kinetochores in Nlrp5-deficient MII oocytes (1.55 ± 0.14, n=21) at a greater intensity than in wildtype oocytes (1.00 ± 0.04, n=16) (p=0.001, Mann-Whitney test) [Figure 17D]. Thus, in the Nlrp5- deficient state, dynactin p50 is retained on kinetochores, which supports our findings that Nlrp5- deficient oocytes have more misaligned MII chromosomes [Figure 12].

Aurora Kinase B A WT Nlrp5KO * 1.3 (p = 0.0260) 1.2 Aurora 1.1 1.0 B 0.9 0.8 DAPI 0.7 0.6 Aurora (RFU) 0.5 0.4 B 0.3

0.2 Chromosomal PlateChromosomal

Relative Expression on Relative Expression 0.1 n=15 n=18 0.0 WT Nlrp5 KO B Blocking WT Nlrp5KO Peptide 1.75 Aurora Kinase C (p = ***0.0001) Aurora 1.50 C 1.25 DAPI 1.00

0.75 (RFU) Aurora 0.50 C

Chromosomal PlateChromosomal 0.25

Relative Expression on Relative Expression n=16 n=20 0.00 WT Nlrp5 KO C WT Nlrp5KO Dynactin p50 ** 1.75 (p = 0.001) Dynactin 1.50 p50 1.25 DAPI 1.00 Dynactin 0.75 p50 0.50 0.25

Relative Expression on Relative Expression n=16 n=21

Chromosomal Chromosomal Plate (RFU) 0.00 WT Nlrp5 KO

Figure 17: Relative expression of various Spindle Assembly Checkpoint proteins in wildtype and Nlrp5-deficient MII oocytes.

Immunocytochemistry was performed in MII oocytes, followed by confocal microscopy focusing on the chromosomal plate, to investigate the protein level of selected proteins. Representative expression patterns and quantification of fluorescence intensity on the metaphase plate. Values represent fluorescence intensity relative to wildtype levels ± SEM. Asterisks denote means are significantly different. Unpaired samples, two-tailed t-tests.

A: In wildtype MII oocytes Aurora B is present in low levels or is almost undetectable and in Nlrp5-deficient oocytes Aurora B localizes to the kinetochores and there is a significant increase in protein expression (p=0.026). Aurora B Antibody (mouse)-green; DAPI-blue.

B: In wildtype oocytes, Aurora C localizes to the kinetochores and in Nlrp5-deficient oocytes this staining is increased, consistent with quantification suggesting that there is a significant increase in Aurora C protein levels in Nlrp5-deficient oocytes (p=0.0001). t-test. Aurora C Antibody (rabbit)-red; DAPI-cyan.

C: In wildtype MII oocytes dynactin p50 localizes to the kinetochores and in Nlrp5-deficient oocytes there is a significant increase (p=0.001). Mann-Whitney Rank Sum test. Dynactin p50 Antibody (mouse)-green; DAPI-blue.

3.2.5 NOCODAZOLE TREATMENT IN METAPHASE II OOCYTES To determine if the disruption of spindles alters native NALP5 expression, we treated wildtype MII oocytes with nocodazole, a spindle-perturbing agent. Nocodazole is a microtubule-depolymerising agent, used to cause the disruption of normal assembly of spindle microtubules.

With nocodazole treatment in MII oocytes, α-tubulin staining showed that spindles were completely disassembled, as evidenced by no intact spindles, while spindles in the vehicle control group (oocytes that were cultured in dimethyl sulfoxide (DMSO)-supplemented medium for 15 minutes) were clearly visible. In the control group, 82% showed strong NALP5 staining on the DNA region, while the other 18% showed only cytoplasmic staining (n=56) [Figure 18]. Upon treatment of nocodazole and spindle disassembly, NALP5 localization to chromosomes decreased compared to DMSO-treated controls (n=9). This data suggest that spindle attachment to chromosomes is required for the maintenance of NALP5 localization to the chromosomal plate and without this attachment NALP5 dissociates or degrades from the plate. This could imply that, in oocytes, the sensing of NALP5 on the chromosomal plate by the spindle signals the proper activation of the SAC and without NALP5 attachment to the spindle, as in the case in Nlrp5-deficient oocytes, the SAC in not appropriately activated.

Vehicle Control (DMSO) Nocodazole 10μg/mL

α-Tubulin

NALP5

Figure 18: NALP5 expression with Nocodazole treatment in MII oocytes.

NALP5 expression after nocodazole treatment in MII oocytes, using laser-scanning confocal microscopy. Wildtype oocytes were treated with nocodazole, spindle depolymerising agent, or DMSO (vehicle control) and immunocytochemistry was performed on oocytes using NALP5 antibody. NALP5 expression decreases when spindle is disrupted. DMSO treatment group n=56; nocodazole treatment group n=9. α-Tubulin (mouse)-green; NALP5 (rabbit)-red.

3.3 THE ROLE OF NLRP5 IN TRIGGERING AN APPROPRIATE DNA DAMAGE

RESPONSE IN OOCYTES

3.3.1 UV IRRADIATION RESPONSE IN MII OOCYTES For our next objective, we wanted to assess whether Nlrp5-deficient oocytes are defective in the hallmarks of DNA damage, as the responses to DNA damage are essential to ensure the transmission of intact genetic information [35]. Recently, it has been shown that zygotes are capable of base excision repair [118, 135]. As already presented, we have observed that Nlrp5- deficient oocytes displayed a higher frequency of possible chromosomal separation, evidenced by scattering of chromosome pieces via immunofluorescence microscopy, suggesting that they may have more damaged DNA.

Initially, we designed an experiment to evaluate whether markers of DNA damage [53BP1, phosphorylated Histone H2AX, phosphorylated Histone H3, BRCA1, and ATM], commonly used in somatic cells, showed differences in expression levels in MII, ovulated oocytes with and without ultraviolet (UV) irradiation, as this has not been previously determined. We induced DNA damage in wildtype MII oocytes, by means of UV exposure and subsequently evaluated the expression levels of these DNA damage response proteins, by performing immunocytochemistry and quantification. We chose to use UV-induced damage as our method of inducing DNA damage because previous publications have reported that oocytes are capable of UV-elicited excision repair or nucleotide excision repair [114]. UV irradiation of somatic cells triggers the DNA-damage response that involves the proteins: ATR, CHK1, γH2AX and 53BP1 [116]. Figure 8 shows the DNA damage pathway in somatic cells. After exposing oocytes to 12 J/m2 of UV radiation, oocytes were cultured for 1 or 5 hours to sense damage and then cells were fixed and immunostained to assess the recruitment of key DNA damage response proteins. Control groups were not UV-exposed, but were cultured for the same amount of time.

Ultimately, we wanted to see whether Nlrp5-deficient oocytes show similar patterns of expression of the DNA damage markers, as observed in wildtype oocytes induced with UV. If so, this could imply that the observed change is due to the DNA damage response in Nlrp5- deficiency.

3.3.1.1 NALP5 AND UV DAMAGE IN MII OOCYTES Firstly, we wanted to assess the effect that UV damage had on NALP5 protein expression, in wildtype oocytes. We observed decreased NALP5 expression on the chromosomal plate and in the cytoplasm, similar to the NALP5 staining pattern evidenced by nocodazole treatment. When we quantified the level of fluorescence of NALP5 on the global (entire) oocyte, we observed that there was a statistically significant decrease of NALP5 expression after UV exposure (19800 ± 750, n=35) compared to control (29800 ± 970, n=21) (p<0.001) [Figure 19]. While there was no significant difference in NALP5 expression between the one hour (24800 ± 850, n=29) and five hours (24900 ± 880, n=27) time points (p=0.991). This suggests that UV radiation causes degradation of NALP5 protein in MII oocytes.

66 Negative 1h Control UV + 1h 5h Control UV + 5h Control

NALP5 DAPI

Global NALP5 ** p<0.001

35000 30000 25000 20000

15000 (RFU) 10000 5000

Fluorescence Intensity Fluorescence n=11 n=18 n=10 n=17 0 1h Control UV + 1h 5h Control UV + 5h

Figure 19: NALP5 expression after UV exposure.

NALP5 expression after UV exposure in MII oocytes, using laser-scanning confocal microscopy. Wildtype oocytes, exposed to UV damage were cultured for 1h or 5h and control groups were not exposed to UV damage, but were cultured for the same time period. After UV treatment, immunocytochemistry was performed on oocytes using NALP5 antibody. Negative control was exposed to secondary antibody alone, donkey anti-rabbit (red). NALP5 (rabbit)-red; DAPI-blue. Quantification of mean fluorescence intensity of the entire oocyte, measured in relative fluorescence units (RFU), corrected against negative control (no primary antibody). There was a significant decrease in NALP5 expression with UV exposure (p<0.001), compared to control groups. Values represent mean fluorescence units ± SEM. Two-way ANOVA, Holm- Sidak method. Asterisks signify mean values between paired groups are significant.

3.3.1.2 PHOSPHORYLATED HISTONE H2AX (ΓH2AX) Histone H2AX is a histone H2A variant [119]. Phosphorylated histone H2AX (γH2AX) at serine 139 is a classical marker of DNA damage and is also present in response to UV irradiation [122]. UV-induced phosphorylation of H2AX requires nucleotide excision repair factors [122].

By conducting our UV treatment in wildtype MII oocytes, we found that there is a statistically significant increase in the fluorescence intensity of γH2AX on the chromosomal plate after UV exposure (12200 ± 1200, n=31) compared to controls (5500 ± 1390, n=24) (p<0.001) [Figure 20A]. Consistent with previous reports, γH2AX expression increases upon UV exposure. Moreover, time of culture after UV exposure affected γH2AX fluorescence levels, because there was a significant increase in γH2AX fluorescence intensity at 5 hours (11700 ± 1300, n=29) than at 1 hour (6000 ± 1300, n=26) after UV (p=0.003), suggesting that at 5 hours γH2AX to able to get recruited to more sites of DNA damage than at 1 hour.

Immunocytochemistry results of γH2AX staining on wildtype and Nlrp5-deficient oocytes MII illustrate that in both genotypes there is localization on the chromosomes, more specifically on the histones, and when quantified Nlrp5-deficient oocytes displayed significantly increased levels of γH2AX accumulation (2.04 ± 0.17, n=11) compared to wildtypes (1.00 ± 0.08, n=17) (p<0.001) [Figure 20B]. One explanation for an increase in γH2AX in Nlrp5-deficient oocytes could be because there may be more H2AX protein, as our microarray data resulted in Nlrp5- deficient oocytes having elevated H2afx (H2ax-related gene) transcripts levels compared to wildtype oocytes [Table 7]. Alternatively, since there is increased H2AX phosphorylation after UV irradiation and because Nlrp5-deficient oocytes have more γH2AX accumulation this could imply that Nlrp5 deficiency causes increased UV damage.

A 1h 5h B Control UV + 1h Control UV + 5h WT Nlrp5KO γH2AX γH2AX

H2AX ** p<0.001

* H2AX p=0.003 ** 2.5 (p<0.001) 20000 2.0 15000 1.5 10000

(RFU) 1.0 (RFU)

5000 0.5

Chromosomal Chromosomal Plate

Relative Expression on Relative Expression Fluorescence Intensity Fluorescence on Chromosomal Plateon n=11 n=15 n=13 n=16 n=17 n=11 0 0.0 1h Control UV + 1h 5h Control UV + 5h WT Nlrp5 KO

Figure 20: γH2AX expression after UV exposure and in Nlrp5 deficiency.

A: Phosphorylated histone H2AX at serine 139 (γH2AX) expression after UV damage in MII oocytes, using laser-scanning confocal microscopy. Wildtype oocytes, exposed to UV damage were cultured for 1h or 5h and control groups were not exposed to UV damage, but were cultured for the same time period. After UV treatment, immunocytochemistry was performed on oocytes using γH2AX antibody. γH2AX (rabbit)-red; DAPI-blue. Quantification of mean fluorescence intensity of the chromosomal plate, measured in relative fluorescence units (RFU). There was a significant increase in γH2AX expression in UV-irradiated groups, compared to control groups (p<0.001). There was also a significant increase γH2AX levels at 5h, compared to 1h (p=0.003). Values represent mean fluorescence units ± SEM. Asterisks signify means between paired groups are significant. Two-way ANOVA, Holm-Sidak method.

B: Phosphorylated histone γH2AX protein expression in wildtype and Nlrp5-deficient MII oocytes. Immunocytochemistry was performed in freshly ovulated MII oocytes to investigate the protein level of γH2AX. Confocal microscopy images reveal increased expression on the histones. γH2AX antibody (rabbit)-red; DAPI-blue. Quantification of fluorescence intensity, measured in relative fluorescence intensity, suggests that there is a significant increase in γH2AX protein expression on the chromosomal plate of Nlrp5-deficient oocytes compared to wildtype oocytes (p<0.001). Values represent mean fluorescence units ± SEM. Asterisks denote values are significant. Mann-Whitney Rank Sum test.

3.3.1.3 53BP1 53BP1 (53-Binding Protein-1) is a DNA damage response factor involved in DNA damage-induced, cell-cycle arrest. When we conducted immunocytochemistry with 53BP1 in wildtype oocytes and quantified the fluorescence intensity of 53BP1 on the chromosomal plate, there was a statistically significant increase after UV exposure (640 ± 20, n=31) compared to control groups (530 ± 20, n=23) (p<0.001) [Figure 21A]. This was due to increased recruitment of 53BP1 in the UV-exposed group, as 6 of 13 oocytes had very obvious staining on the chromosomes, which was absent in the control group (n=10). Furthermore, the increase of 53BP1 after UV is evenly distributed on the chromosomes in MII oocytes, as opposed to being present at foci, consistent with a previous report in fibroblast cells [122]. This indicates that UV exposure in wildtype MII oocytes causes the increase of 53BP1 on the chromosomal plate.

When wildtype and Nlrp5-deficient oocytes were stained with the same antibody, we observed that the wildtypes had a low level of 53BP1 accumulation on chromosomes that was distributed quite evenly. In Nlrp5-deficient oocytes (1.92 ± 0.11, n=14), the fluorescence intensity of 53BP1 on the chromosomal plate was significantly increased compared to wildtypes (1.00 ± 0.04, n=14) (p<0.001) [Figure 21B]. Taken together, increased 53BP1 expression after UV exposure and increased 53BP1 expression in Nlrp5-deficient oocytes suggest that Nlrp5 deficiency could lead to increased DNA damage.

1h 5h A B Control UV + 1h Control UV + 5h WT Nlrp5 KO

53BP1 53BP1 DAPI DAPI

53BP1 ** p<0.001

53BP1 750 ** 2.25 (p<0.001) 2.00 1.75 500 1.50 1.25

(RFU) 1.00 250 (RFU) 0.75 0.50

Chromosomal Chromosomal Plate 0.25 Fluorescence Intensity Fluorescence on Chromosomal Plateon n=10 n=13 n=13 n=18 Relative Expression on Relative Expression n=14 n=14 0 0.00 1h Control UV + 1h 5h Control UV + 5h WT Nlrp5 KO

Figure 21: 53BP1 expression after UV exposure and in Nlrp5 deficiency.

A: 53BP1 expression after UV damage in MII oocytes, using laser-scanning confocal microscopy. Wildtype oocytes, exposed to UV damage were cultured for 1h or 5h and control groups were not exposed to UV damage, but were cultured for the same time period. After UV treatment, immunocytochemistry was performed on oocytes using γH2AX antibody. 53BP1 (rabbit)-red; DAPI-blue. Quantification of mean fluorescence intensity of the chromosomal plate, measured in relative fluorescence units (RFU). There was a significant increase in 53BP1 expression with UV exposure (p<0.001), compared to control groups. Values represent mean fluorescence units ± SEM. Two-way ANOVA, Holm-Sidak method. Asterisks signify means between paired groups are significant.

B: 53BP1 protein expression in wildtype and Nlrp5-deficient MII oocytes. Immunocytochemistry was performed in MII oocytes to investigate the protein level of 53BP1. 53BP1 antibody (rabbit)-red; DAPI-blue. Quantification of fluorescence intensity, measured in relative fluorescence intensity, suggests that there is a significant increase in 53BP1 protein levels on the chromosomal plate of Nlrp5-deficient oocytes compared to wildtype oocytes (p<0.001). Values represent mean fluorescence units ± SEM. Asterisks denote values are significant. Mann-Whitney Rank Sum test.

3.3.1.4 PHOSPHORYLATED HISTONE H3 AT SERINE 10 (H3S10) Phosphorylation of histone H3 at serine 10 is strongly correlated with chromosome condensation during mitosis and meiosis [125]. Quantification of the fluorescence intensity of phosphorylated H3 on the chromosomal plate in wildtype oocytes revealed that there was no significant difference of H3S10 accumulation between UV-exposed groups (1300 ± 90, n=39) and non-exposed groups (1400 ± 130, n=23) (p=0.564) [Figure 22A]. There was a statistically significant increase in the fluorescence intensity of H3S10 in the five hours of incubation groups (1800 ± 100, n=35) compared to the one hour of incubation groups (860 ± 130, n=27) (p<0.001). These results indicate that prolonged culturing or post-ovulatory aging, rather than UV exposure, increases the accumulation of phosphorylated histone H3.

Additionally, we found that there was a trend towards an increase in the fluorescence intensity of phosphorylated histone H3 at serine 10 on the chromosomal plate in Nlrp5-deficient oocytes (1.29 ± 0.12, n=20), compared to wildtype oocytes (1.00 ± 0.09, n=26), although this was not significant (p>0.05) [Figure 22B]. Since we saw that H3S10 accumulation increases with 5 hours of culture in wildtype oocytes, taken together, these results imply that Nlrp5 deficiency causes the H3 histone status to appear like that of aged oocytes. This data is consistent with our Aurora kinases protein expression data, in that there were increases in both Aurora kinases B and C expression in Nlrp5-deficient oocytes, which phosphorylate histone H3 at serine 10 [Figure 17A-C]. Alternatively, serine 10 of histone H3 is a substrate for purified phosphoprotein phosphatase 1 (PPP1C) during meiosis I [73] and our results could suggest that Nlrp5-deficient oocytes may have decreased levels of PPP1C. Furthermore, in accordance with Wei and group, it could be suggested that Nlrp5-deficient oocytes undergo abnormal chromosomal condensation, resulting in the inability to ensure fidelity of chromosome segregation, causing breakage of genetic material [136].

A 1h 5h B Control UV + 1h Control UV + 5h WT Nlrp5 KO H3S10 H3S10

H3S10 ** p<0.001 H3S10 2500 p=0.078 1.5 2000

1500 1.0

(RFU) 1000 (RFU) 0.5

500

Chromosomal Chromosomal Plate Fluorescence Intensity Fluorescence on Chromosomal on Plate n=7 n=20 n=16 n=19 Relative Expression on Relative Expression n=26 n=20 0 0.0 1h Control UV + 1h 5h Control UV + 5h WT Nlrp5 KO

Figure 22: H3S10 expression after UV exposure and in Nlrp5 deficiency.

A: Phosphorylated histone H3 at serine 10 (H3S10) expression after UV exposure in MII oocytes, using laser-scanning confocal microscopy. Wildtype oocytes, exposed to UV damage were cultured for 1h or 5h and control groups were not exposed to UV damage, but were cultured for the same time period. After UV treatment, immunocytochemistry was performed on oocytes using H3S10 antibody. H3S10 (rabbit)-red; DAPI-blue. Quantification of mean fluorescence intensity on the chromosomal plate, measured in relative fluorescence units (RFU). There was a significant increase in H3S10 levels at 5h compared to 1h (p<0.001). Values represent mean fluorescence units ± SEM. Asterisks signify means between paired groups are significant. Two-way ANOVA, Holm-Sidak method.

B: Phosphorylated histone H3 at serine 10 (H3S10) protein expression in wildtype and Nlrp5- deficient MII oocytes. Immunocytochemistry was performed in freshly ovulated MII oocytes to investigate the protein level of H3S10. Confocal microscopy images reveal increased expression on the histones. H3S10 antibody (rabbit)-red; DAPI-blue. Quantification of fluorescence intensity, measured in relative fluorescence intensity, suggests that there is an increase in H3S10 protein expression on the chromosomal plate of Nlrp5-deficient oocytes compared to wildtype oocytes (p>0.05). Values represent fluorescence intensity relative to wildtype levels ± SEM. Mann-Whitney Rank Sum test.

3.3.1.5 BRCA1 BRCA1 has been shown to be needed for meiotic spindle assembly and SAC activation in mouse oocytes since its depletion causes misaligned chromosomes and impaired spindles [106]. Additionally, BRCA1 has an established role in DNA damage signalling and repair [126, 137].

By conducting our UV treatment in wildtype MII oocytes, we found that BRCA1 was expressed on the spindle poles in UV-exposed and control groups [Figure 23A]. Quantification of BRCA1on the spindle poles, followed by statistical analysis, revealed that there is a statistically significant interaction between UV exposure and the time of culture after UV exposure (p<0.001), suggesting that the effect of UV exposure depends on culture period. We also observed that at both 1 hour (1300 ± 60, n=9) and 5 hours (1200 ± 60, n=6) after UV irradiation, there was a statistically significant increase in BRCA1 fluorescence intensity on the spindle poles compared to the respective controls (1h (500 ± 50, n=9) (p<0.001) and 5h (800 ± 50, n=12) (p<0.001), respectively). This suggests that BRCA1 expression increases with UV- induced damage in MII oocytes. There was also a significant increase in BRCA1 expression on the spindle poles in the five hour control compared to the one hour control (p<0.001), implying that BRCA1 could be unregulated by post-ovulatory aging. After UV irradiation, BRCA1 localizes to the spindle poles and does not become nuclear. This would indicate that the dose of UV may also cause spindle damage and this contributes to increased BRCA1 accumulation on the spindle.

Immunocytochemistry results of BRCA1 staining on wildtype and Nlrp5-deficient MII oocytes illustrate that in both genotypes there is localization on the spindle poles and when quantified Nlrp5-deficient oocytes (1.13 ± 0.04, n=18) displayed significantly increased levels of BRCA1 compared to wildtypes (1.00 ± 0.04, n=21) (p=0.0224, t-test) [Figure 23B]. These results also imply that abnormal spindle function caused by Nlrp5 deficiency causes increased BRCA1 recruitment to the spindle.

A 1h 5h B Control UV + 1h Control UV + 5h WT Nlrp5 KO BRCA1 BRCA1 DAPI DAPI

BRCA1 BRCA1 1500 b * 1.2 (p=0.0224) b 1.1 1.0 1000 0.9 c 0.8 0.7 a 0.6 500 0.5 0.4 0.3

0.2

Spindle Poles (RFU) Poles Spindle Fluorescence Intensity Fluorescence

on Spindle Poles (RFU) Spindle Poles on n=9 n=9 n=12 n=6 Relative Expression on Relative Expression 0.1 n=21 n=18 0 0.0 1h Control UV + 1h 5h Control UV + 5h WT Nlrp5 KO

Figure 23: BRCA1 expression after UV exposure and in Nlrp5 deficiency.

A: BRCA1 protein expression after UV exposure in MII oocytes, using laser-scanning confocal microscopy. Wildtype oocytes, exposed to UV damage were cultured for 1h or 5h and control groups were not exposed to UV damage, but were cultured for the same time period. After UV treatment, ICCH was performed on oocytes using a BRCA1 antibody. BRCA1 (1904) antibody (rabbit)-red; DAPI-blue. Quantification of mean fluorescence intensity on the spindle poles, measured in relative fluorescence units (RFU). BRCA1 is affected by UV exposure, evidenced by accumulation on the spindle poles, depending on the time period of incubation. There is a statistically significant interaction between UV exposure and the time points (p<0.001). Values represent mean fluorescence units ± SEM. Different letters above bars signify means between bars are significant. Two-way ANOVA, Holm-Sidak method.

B: BRCA1 protein expression in wildtype and Nlrp5-deficient MII oocytes. Immunocytochemistry was performed in freshly ovulated MII oocytes to investigate the protein level of BRCA1. Confocal microscopy images reveal increased expression on the spindle poles. BRCA1 (1904) antibody (rabbit)-red; DAPI-blue. Quantification of fluorescence intensity, measured in relative fluorescence intensity, suggests that there is a significant increase in BRCA1 protein expression on the spindle poles of Nlrp5-deficient oocytes compared to wildtype oocytes (p=0.0224). Values represent fluorescence intensity relative to wildtypes ± SEM. Asterisks denote values are significant. Unpaired samples, two-tailed t-tests.

3.3.1.6 ATM We also analysed the phosphorylation status of the ATM protein kinase (Serine 1981), a site known to be activated in response to DNA damage, but did not find a difference in staining pattern and expression between UV-exposed and non-exposed MII oocytes. Staining was cytoplasmic with and without UV exposure and there was no significant change in fluorescence intensity.

3.3.2 COMET ASSAY IN MII OOCYTES WITH NLRP5 DEFICIENCY The comet assay is a single cell gel electrophoresis assay used to assess DNA damage in cells by detecting single-stranded DNA breaks, double-stranded breaks, and may detect a few apurinic and apyrimidinic sites. The assay is based on the premise that denatured and cleaved DNA fragments are able to migrate faster due to an electric current and will create a comet-like halo, whereas undamaged DNA will remain in place with no or minimal migration.

When we performed immunocytochemistry on wildtype oocytes and exposed them to UV irradiation, we observed that there was an increased frequency of chromosomes that were scattered and misaligned compared to non-UV-exposed oocytes, which had aligned chromosomes at the metaphase plate [Figure 24A]. To assess if the expression pattern of the DNA damage markers was caused by DNA damage in Nlrp5-deficient oocytes, we performed the comet assay. The control group had MII oocytes that were wildtype and were not exposed to UV [Figure 24B]. These oocytes had DNA that appeared to remain in one place, as evidenced by no halos and nominal-sized comet tails. Doxorubicin was used as a positive control, since it is a chemical that induces both single and double stranded DNA breaks in oocytes by intercalating with DNA [99]. With Doxorubicin treatment, we saw typical-structured comets; where a halo was present near the tail. With UV exposure, we saw the presence of atypical comet structures, having tails that were more dispersed, with a weak diffuse signal. This pattern could be correlated to the observed higher number of scattered chromosomes by UV, evidenced by DAPI staining [Figure 24A]. We found that wildtype and Nlrp5-deficient metaphase-II- arrested oocytes had comparable DNA spots (Nlrp5-deficient (n=12); wildtype (n=7)), since both sets had DNA with nominal comets [Figure 24C], similar to controls. These results indicate that Nlrp5-deficient oocytes do not exhibit DNA damage or these results could indicate that the DNA damage extent is below the threshold of damage detection by the comet assay in metaphase II-arrested oocytes.

A DAPI B C WT

Control Control

Doxorubicin Nlrp5 KO

PB UV UV

Figure 24: Comet assay, measuring DNA damage, in MII oocytes.

A: Chromosomal staining of non-UV-exposed control MIIs showing aligned chromosomes at the metaphase plate, while the UV-exposed chromosomes appear dispersed and scattered. DAPI-blue.

B: Comet assay, measuring DNA damage, was performed in wildtype MII oocytes, using laser- scanning confocal microscopy. Control group displayed nominal-sized comet tails, while the Doxorubicin-exposed oocytes had classical-structured comets, with halo-tails. (Arrowheads denote the head of the comet; arrows denote the tail). The UV-exposed group has comets that are more dispersed with a diffuse signal, reflecting the dispersed nature of the scattered chromosomes. Stained with SYBR Safe Dye.

C: DNA damage in Nlrp5 deficiency. Comet assay was performed in WT and Nlrp5-deficient (KO) MII oocytes. Both wildtype (n=7) and Nlrp5-deficient (n=12) MIIs displayed nominal- sized comet tails, suggesting that DNA damage is not present in Nlrp5-deficient MII oocytes. Stained with SYBR Safe Dye. PB-polar body.

3.3.3 UV DAMAGE AND Α-TUBULIN Immunocytochemistry was performed on wildtype, MII oocytes exposed to UV irradiation (n=37) and non-UV-exposed oocytes (n=17), using α-tubulin to analyse spindle morphology after UV exposure [Figure 25]. Non-UV-irradiated control oocytes showed strong spindle staining with undetectable or low cytoplasmic staining, whereas UV-exposed oocytes displayed fainter spindles with increased cytoplasmic staining at 30 minutes, 1 hour and 5 hours after UV exposure. Furthermore, we also found that many oocytes had spindles that were abnormal after UV irradiation (21/27) compared to non-irradiated controls (0/20). Many of the abnormal spindles were disorganized with multiple poles or had detached microtubules and scattered chromosomes. This data implies that UV irradiation causes spindle damage and the difference in expression of DNA damage markers in Nlrp5 deficiency could be due to UV irradiation causing spindle damage.

Control

α-Tubulin DAPI

UV-exposed

α-Tubulin DAPI

Figure 25: α-Tubulin localization after UV exposure.

Immunocytochemistry with α-tubulin in MII oocytes after UV exposure, using laser-scanning confocal microscopy. Wildtype oocytes were exposed to UV irradiation and ICCH was performed on oocytes using α-tubulin antibody. At 1h and 5h after UV exposure, UV-exposed MII oocytes had fainter spindles, higher cytoplasmic background (indicating partial depolymerisation of spindles) staining and had more spindles of abnormal morphology (37/37) compared to non-exposed controls (4/17). α-tubulin (mouse)-green; DAPI-blue.

3.3.4 DNA DAMAGE MARKERS AND NOCODAZOLE Since we did not observe DNA damage in Nlrp5-deficient oocytes by the Comet Assay, we wanted to see whether the increased levels of DNA damage markers, γH2AX and 53BP1, could be caused by increased spindle damage. To analyze whether spindle depolymerisation causes a difference in γH2AX or 53BP1 expression on chromosomes, we treated MII oocytes with nocodazole and performed immunocytochemistry. Nocodazole treatment did not change the γH2AX expression between vehicle (DMSO) (11720 ± 1050 RFU, n=17) and nocodazole- treated (11450 ± 1320 RFU, n=11) groups (p>0.05) [Figure 26]. Similarly with nocodazole treatment, 53BP1 expression was not altered between vehicle control (11290 ± 450 RFU, n=9) and nocodazole (10180 ± 360 RFU, n=20) groups (p>0.05). These results indicate that spindle damage, caused by depolymerisation of spindles, does not affect the expression of DNA damage markers, 53BP1 and γH2AX. In addition, the upregulation of these markers observed after UV exposure was in fact caused by the increased response to DNA damage and not due to the effects of spindle damage. Taken altogether, this data implies that DNA damage may be present in Nlrp5-deficient oocytes; however the comet assay may not be sensitive enough to detect it.

A DMSO Noc B DMSO Noc 53BP1 γH2AX DAPI 53BP1

53BP1 H2AX 12000 13000 11000 12000 10000 11000 9000 10000 8000 9000 7000 8000 7000 6000 6000 5000 5000 4000 4000 3000 3000 2000 2000 1000 n=9 n=20 1000 n=17 n=11

0 0

Mean Fluorescence Intensity Fluorescence Mean

Mean Fluorescence Intensity Fluorescence Mean on Chromosomal on Plate (RFU) on Chromosomal on Plate (RFU) DMSO Nocodazole DMSO Nocodazole

p>0.05 p>0.05

Figure 26: 53BP1 and γH2AX expression after Nocodazole treatment.

Wildtype MII oocytes, exposed to nocodazole (Noc) treatment, followed by indirect immunocytochemistry using 53BP1 and γH2AX antibodies. Quantification of mean fluorescence intensity of the chromosomal plate, measured in relative fluorescence units (RFU) ± SEM. There is no statistically significant change in the expression of 53BP1 and γH2AX with nocodazole treatment. Unpaired samples, two-tailed t-tests. A: 53BP1 (rabbit)-red; DAPI-blue. B: γH2AX (rabbit)-red.

3.4 MICROARRAY RESULTS We performed a microarray study using wildtype and Nlrp5-deficient metaphase II- arrested oocyte samples to investigate what pathways are deregulated due to Nlrp5 deficiency. Table 6 and Table 7 show a list of selected microarray targets with relevance to the Spindle Assembly Checkpoint or DNA damage response. Fold changes are based on 4 sets of datasets and the fold difference between wildtype and Nlrp5-deficient MII oocytes are listed below. Two fold differences per gene represent two different probe sets.

For a complete list of all gene targets that were significantly different between wildtype and Nlrp5-deficient oocytes, please see Tables 9 and 10 in the appendix section.

Table 6: mRNA transcripts with Decreased Expression in Nlrp5-Deficient MII oocytes Gene Gene Name Fold t-test Notes Difference (n=4) Nlrp5 NLR family, 8.21 p<0.001 - highest fold difference among analysed genes pyrin domain containing 5 Cdkl2 cyclin- 2.58 p=0.089 - Cdc2-related serine/threonine protein kinase dependent 2.34 p=0.05 kinase-like 2 (CDC2-related kinase) Kif3b kinesin family 2.13 p<0.01 - MT binding motor protein member 3B - Kif3b is down-regulated in aged oocytes [57] Mcl1 myeloid cell 2.11 p<0.01 - Mcl1 KO show spindles defects leukemia 2.01 p=0.01 sequence 1 Mtap9 microtubule- 1.93 p<0.05 - Involved in mitosis, cell cycle, and cell associated division protein 9 AurKC Aurora kinase 1.76 p<0.0001 - germ cell-specific Aurora kinase C

Table 7: mRNA transcripts with Increased Expression in Nlrp5-Deficient MII oocytes Gene Gene Name Fold t-test Notes Differenc (n=4) e Dync1i2 dynein 3.87 p<0.005 - Cytoskeletal protein cytoplasmic 1 - Dynein, cytoplasmic, light polypeptide 2B is intermediate over-expressed in human aneuploid oocytes chain 2 [138] H2afx H2A histone 3.14 p<0.01 - Positive regulation of DNA repair family, - H2A activity is modulated by Bub1 [139] member X

4 LIMITATIONS

For all of the analysis of spindle checkpoint protein expression and UV damage marker expression, only normal, healthy-appearing metaphase II oocytes (about 80-90% of entire population) were used for quantification. In particular, with regards to spindle morphology evaluation, our analysis was limited to the selection of the samples that appeared to have normally aligned chromosomes and spindles that were suitably oriented (i.e., longitudinal orientation). Malformed spindles were omitted from quantitative analysis; however were deemed important for qualitative analysis. This was done to ensure that the scored values were not inaccurately skewed.

Furthermore for Nlrp5 and NALP5 analysis, our experiments were limited to the assessment of the gene and protein in mouse oocytes. Unfortunately, cell lines were not used as model as to investigate Nlrp5 function, as no oocyte-specific cell lines are available. In addition, no other cells, aside from sperm cells, undergo meiosis and thus there may be a unique apparatus in place that regulates this. Additionally, NALP5 has been shown to have interacting partners, many of which are also oocyte-specific, hence they will also not be present in regular cells and consequently will not be a good representative model of an oocyte. As mentioned earlier, in section 1.6.4, the Nlrp5-deficient (or knockout) mouse model is a useful model to study infertility; however, along with this comes a few limitations. Each female mouse only ovulates about 15 to 20 oocytes; hence there is a limited supply of samples. In addition, many assays require plenty of oocytes, such as Western blots, ELISA assays or co- immunoprecipitation assays. For example, to perform a Western blot, about 100-500 oocytes per genotype or treatment would be needed to assess one protein. Moreover, the breeding colony performance is not always predictable, irrespective of the genotype or strain of mouse. Furthermore, female mice do not always ovulate and sometimes do not release a sufficient number of oocytes.

Ultimately, it would be interesting to evaluate NALP5 expression in human oocytes or embryos allowing for the comparison between gametes from fertile and infertile patients. Unfortunately with human samples we are limited to not having a reliable control, as fertile women will not undergo assisted reproductive treatment to retrieve their oocytes. Typically, human samples are more readily available from infertile or subfertile patients who seek assisted reproduction treatment and undergo gamete retrieval. The best control oocytes that can be used

82 are from patients who seek assisted reproductive treatment due to male factor infertility or patients who have tubal (reproductive tract) factor infertility.

As antibodies recognizing spindle assembly checkpoint proteins have previously been tested in oocytes for immunofluorescence analysis, I conducted my experiments in accordance with previous protocols using the same antibodies. However, the same antibodies did not work on oocytes when I tried both Formalin and PHEM fixation. Many of these antibodies that were designed to localize to kinetochores did not work in whole-mount oocyte immunocytochemistry procedures. As a result, I also performed another assay called chromosomal spreads, followed by immunocytochemistry. The premise of this assay is to cause the cytoplasm of the egg to disintegrate, thereby allowing for the fixation of vastly spread out chromosomes. This thereby allows for more access of the antibody to specific regions of the chromosomes. Unfortunately, the conditions for this technique were not fully optimized and thus resulted in incomplete spreading of chromosomes. Table 8 depicts of list of antibodies which did not result in specific signal by immunofluorescence. In correspondence with authors that used the same antibodies in publications, they mentioned that the antibody performance could depend on the batch or lot number and they have also had difficulty repeating some of their published work.

Table 8: List of Antibodies that did not work

Antibody Company Catalogue # purified Mouse Anti-IAK1; Aurora A Tranduction lab I71320 mouse monoclonal -Bub1; 1mg/ml Chemicon - goat polyclonal -Bub1 Santa Cruz sc-18286 goat polyclonal -Bub3 Santa Cruz sc-19395 mouse -BubR1 MAB3612 Chemicon - human-ANA-Centromere Autoab = Crest Cortex CS1058 human -CENP-E Aliquot - human -Crest Aliquot - mouse -Hec1 Santa Cruz sc-135934 purified Mouse Anti-MAD2; anti-hsMAD2 mAb Tranduction lab H57520

5 DISCUSSION

A plethora of factors can contribute to the cause of infertility. Of particular interest to my work is a focus on infertility in couples who have repeated assisted reproductive cycle failure, which partially stems from the embryo having a compromised developmental potential. This in turn, can be attributed to poor quality oocytes produced by the female. Although it has been well understood that the quality of the oocyte is a prognostic factor in determining a healthy pregnancy, very little is known about the molecular mechanisms underlying differences in developmental competence of an oocyte. Maternal-effect genes, over the past ten years, have been shown to be crucial for normal embryonic development in mammalian models. Many of these genes are in fact oocyte- or preimplantation embryo-specific, with a unique pattern of expression. Previous studies using mammalian knockout models of specific maternal-effect genes caused early embryo loss followed by infertility [45-47]. Another molecular mechanism that influences the quality of all cells, including the oocyte, is genomic stability. Since the oocyte is the progenitor of the zygote, which eventually becomes an embryo and foetus, it is crucial that the founding oocyte maintains its genomic integrity. Defects in the maintenance of genome stability have been shown to result from deficiencies in the surveillance mechanisms that are linked to proper segregation of DNA, such as the spindle assembly checkpoint, and the surveillance mechanisms that lead to DNA repair. Consequences of genomic instability include: chromosomal segregation errors, which result in multinucleation and can ultimately progress to aneuploidy. Aneuploidy has been well correlated with maternal age, as older women (greater than 40 years) seeking in vitro fertilization treatment produce 50% of oocytes that are chromosomally abnormal [140]. Although a fraction of aneuploid embryos implant and produce children with congenital disorders such as, Down’s syndrome, the vast majority of these embryos fail to implant or culminate in spontaneous abortions. This overview proposes that investigating genome integrity in oocytes will help elucidate information about the molecular mechanism of oocyte quality.

In this study, we have described exciting new phenotypes that could possibly explain a new set of outcomes in female infertility. It has been established that female infertility in mice and humans is linked to aneuploidy, which in turn has been commonly correlated with maternal aging [12, 14-16, 141]. Furthermore, aside from Nlrp5 being expressed in human oocytes, there is limited information regarding the role of this gene in female infertility. Since Nlrp5 transcript

84 expression decreases with aging [57] and since aging is associated with the high incidence of aneuploidies, this fits into the aneuploidy/infertility puzzle. Additionally, spindle abnormalities and faulty chromosome alignment on the metaphase plate are associated with advanced maternal age and likely influence the increased frequency of aneuploidy with age [77]. In our study, we have found that Nlrp5-deficient metaphase-II arrested (MII) oocytes display a higher frequency of spindle morphology defects. It was recently reported that about 4% of young mouse MII oocytes had abnormal chromosomal alignment, whereas about 18% of displaced/misaligned chromosomes were seen in old oocytes [16]. Likewise, in our study when analyzing chromosomal alignment abnormalities, we found that in Nlrp5-deficient oocytes there was an incidence of 22%, while there was a 3% defect incidence in wildtypes. These findings highlight that the frequency of chromosomal alignment is strikingly similar between Nlrp5-deficient and old, ovulated oocytes. Based on our study, we have also found that zygotes derived from Nlrp5- deficient mouse oocytes have increased rates of multinucleation. The extra pronuclei that were present in polyploid zygotes were of maternal origin, implying that there were likely meiotic deficiencies in the oocyte that preceded these outcomes in the zygotes. These findings are strengthened by human studies in that human infertility has been associated with multinucleation in embryos [18, 19]. These data suggest that Nlrp5 or downstream molecular pathways affected by Nlrp5 deficiency may potentially be responsible for multinucleation that is evident in human cases.

An important NALP5-interacting partner is FILIA, which is encoded by the maternal- effect gene, Filia. Interestingly, studies have demonstrated that Filia knockout mice display a spectrum of phenotypes similar to Nlrp5-deficient mice [46]. Since Filia knockouts exhibit these phenotypes in the two- to four-cell embryo stages and because the absence of Filia does not affect meiosis I or II, as evidenced by normal spindle morphology, Nlrp5 seems to play a role earlier on in development at the ovulated oocyte stage. Filia knockout females are subfertile because they produce smaller litters, indicating that disruption of this gene produces a milder phenotype compared to Nlrp5. Exhibiting a less dramatic phenotype implies that Filia likely works downstream of Nlrp5. While FILIA localizes subcortically in oocytes and embryos to the subcortical maternal complex, its involvement in the Spindle Assembly Checkpoint (SAC) and how it results in aneuploidy is currently unclear [47]. Normally in interphase FILIA has a subcortical distribution; however its localization has not been explored during mitosis.

Alternatively, even if FILIA does not localize to chromosomes during mitosis, it could have secondary effects by causing post-translational modification and or activation of SAC proteins.

We observed that the use of different fixatives can impact the visualization of protein localization. Previous reports have shown that NALP5 protein is expressed in the subcortex of metaphase II-arrested oocytes, when fixed in 1-2% paraformaldehyde [47, 64]. We have also observed this pattern with formalin fixation (data not shown). However, by using PHEM fixative we saw that the NALP5 protein is cytoplasmic with enrichment on chromosomes in MII oocytes, more specifically on the kinetochores, which is commonly where SAC-involved proteins are expressed. PHEM fixative contains a detergent that preferentially retains proteins that are permanently bound to the cytoskeleton, while removing soluble structures yielding lower cytoplasmic staining compared to other fixatives. We believe the NALP5 staining pattern observed in ovulated oocytes is real because the fluorescence intensity of NALP5 is dramatically reduced in Nlrp5-deficient age-matched-oocytes. Additionally, in the Filia knockout mouse model, knockout embryos shared similar SAC-impaired phenotypes compared to Nlrp5 oocytes, but did not have an explanation as to how FILIA’s protein localization to the subcortical maternal complex could influence the SAC. Recently, it has been reported that NALP5 confocal immunofluorescence staining with cortical enrichment depends on conditions of fixation [48]. This is consistent for other cortically enriched proteins as fixation and immunostaining conditions will affect signal intensity [142].

The localization of NALP5 can be possibly explained by its leucine rich repeat domain. As these repeats form a domain involved in protein-protein interaction, this domain may cause NALP5 to interact with another protein that localizes to kinetochores. In accordance, NALP5- interacting member, TLE6, has a nuclear localization signal, which could function in causing the recruitment of NALP5 to the nuclear pores in germinal vesicles, as this expression pattern has been previously reported [64]. Moreover, NALP9 has been found on spindle-chromosome complexes during proteomic mining of these structures in mouse MII oocytes [143].

5.1 SPINDLE ASSEMBLY CHECKPOINT Another contributing factor to the increased incidence of aneuploidy that is associated with maternal age could be the reduced strength of the SAC in old oocytes, since chromosomes will separate at anaphase even if proper kinetochore-microtubule attachment has not been established. This is further supported by the gene expression of old human oocytes, as these oocytes display dysregulation of transcript accumulation of Spindle Assembly Checkpoint (SAC) components [16, 17].

In Nlrp5-deficient MII oocytes the SAC proteins BUB1 and BUBR1 showed decreased protein expression and did not appear to localize correctly to kinetochores, in contrast to wildtype oocytes. Typically when BUB1, BUBR1, TAp73 are present and the SAC is active, as in wildtype oocytes, if there is an unattached kinetochore, BUB1, BUBR1 and other proteins would activate the SAC, by pausing meiosis at metaphase, and only once the kinetochore properly attaches to the microtubule, meiosis will continue. However, in the case of Nlrp5- deficient oocytes with decreased BUB1, BUBR1, and TAp73, if there is an unattached kinetochore, the SAC appears to not get activated and instead of delaying meiosis until the kinetochore gets attached to the microtubule, the unattached kinetochore and its associated spindle will continue on in meiosis, resulting in improper segregation of chromosomes causing multinucleation. Along the same lines, in Filia knockout embryos, there was decreased protein expression of SAC regulators: MAD2, Aurora kinase A, and Polo-like kinase [46]. Contrarily, in somatic cells, it has been well-reported that SAC proteins such as BUB1, BUBR1, MAD1 and MAD2 localize to misaligned chromosomes [81, 83-85]. However, in Nlrp5-deficient mature oocytes, we see decreased expression of BUB1 and BUBR1 compared to wildtype oocytes. Additionally, we observed that the transactivation domain-containing p73 isoform, TAp73, was decreased from the spindle in Nlrp5-deficient oocytes, which is consistent with previous TAp73 reports in oocytes indicating that decreases in TAp73 result in decreased tethering of BUB1 and BUBR1 to kinetochores [91]. This expression pattern of SAC proteins is temporally-regulated as MII oocytes show lower expression levels of these proteins, in comparison to MI oocytes. It was recently shown that shortly after germinal vesicle break down, BUB1 localized to chromosomes, while in ovulated oocytes BUB1 failed to localize to the chromosomes and was only cytoplasmic [91]. Taken together, this data suggests that several SAC components, along with Nlrp5, are needed for the proper localization of the whole complex of SAC proteins in mature oocytes.

Consistent with previous reports of Aurora kinase B localization in MII oocytes, we observed low to undetectable levels of endogenous protein in wildtype oocytes [92]. A study looking at the SAC in somatic cells investigated the two pathways involved in the maintenance of BUBR1-mediated inhibition of APC/CCdc20 [84]. In the first pathway BUB1 functions as a sensor for improper kinetochore to microtubule attachment and in the second pathway Aurora B functions in maintaining kinetochore biorientation, both of these pathways then lead to the activation of the SAC [84]. In Nlrp5-deficient oocytes, since we observed decreased BUB1 expression and increased Aurora B expression, if the SAC does get activated properly, the checkpoint may potentially be activated by the Aurora kinase B arm, causing the SAC to be more stringent on the biorientation of kinetochores, rather than typical kinetochore to microtubule attachment. These results imply that although there is dysregulation of SAC components with Nlrp5 deficiency, the SAC may still actually get activated by this alternate pathway in a subset of oocytes, as we only see 28% of spindle morphology abnormalities in oocytes and 35% of multinucleation in zygotes, indicating that the pathway correctly responded in the remaining oocytes and zygotes.

The following Aurora kinase that we investigated was Aurora C. In our microarray analysis, although the gene targets have not been verified, in Nlrp5-deficient ovulated oocytes we observed a decrease of Aurora kinase C transcript levels and in our immunocytochemistry experiments we observed an increase in Aurora C protein levels. Since MII oocytes are transcriptionally incompetent, the observed decrease in mRNA can reflect its translational recruitment in order to produce more protein. Mature, ovulated oocytes are transcriptionally inactive due to condensed chromatin structure that denies access to transcriptional machinery; hence less Aurora C mRNA could mean increased utilization of mRNA to make more protein. This pattern of expression concurs with other findings, in MII oocytes that were treated with doxorubicin, there was decreased Bax mRNA because it was used to produce more BAX protein [144]. Another reason for decreased Aurora C mRNA could be due to increased degradation of mRNA in Nlrp5-deficient oocytes. However, decreased amount of mRNA does not always mean increased protein levels. Also in oocytes, Brca1 transcript levels decreased with increased age and resulted in lower BRCA1 protein expression compared to the expression in young oocytes [16]. Taken together, these findings highlight that there is a dynamic mRNA and protein expression pattern in oocytes and that this pattern is dependent on the gene/protein.

The final SAC protein that we investigated in response to Nlrp5 deficiency was dynactin p50. Since dynactin p50 is needed for the removal of MAD1, MAD2 and BUBR1 from aligned kinetochores, perhaps this could also explain why we observe a decrease in BUBR1 levels in Nlrp5-deficient oocytes. Increased levels of dynactin p50 may remove BUBR1 even from unaligned kinetochores [77]. Moreover, overexpression of dynactin p50 in COS-7 cell lines has been previously reported to result in perturbations in mitotic progression [101, 145] partially due to chromosomal misalignment. This report also described that the majority of spindles had two- half spindles that were distorted by having pronounced asymmetry in size, shape, microtubule density and were oriented independently. These results indicate that too much retention of dynactin p50 does not yield normal chromosomal alignment nor spindle morphology characteristics. Furthermore, from our microarray studies, although this has not yet been confirmed by qRT-PCR; we observed that dynein, cytoplasmic 1, intermediate chain 2 (Dync1i2) transcripts were significantly elevated in Nlrp5-deficient oocytes. Interestingly, dynein, cytoplasmic, light polypeptide 2B (DNCL2B), another member of dynein complex, has been found to be overexpressed in human aneuploid oocytes [138]. These findings indicate that Nlrp5-deficient oocytes share decreased dynein transcript levels, similarly with aneuploid human oocytes.

The BRCA1 protein has countless roles that span from spindle assembly in meiotic oocytes to DNA damage repair in somatic cells. We observed that upon UV-induced damage, there was increased accumulation of BRCA1 on the spindle poles and was not nuclear as anticipated, suggesting that the significant increase of BRCA1 expression that was found in Nlrp5-deficient oocytes could indicate spindle damage and not DNA damage. Previous reports in somatic cells using anti-mitotic drugs, such as nocodazole, have correlated BRCA1 with spindle damage [126] and BRCA1 has been postulated to function in preserving genome integrity in meiosis by regulating the spindle assembly checkpoint in mouse oocytes [106]. More specifically, in meiosis BRCA1 recruits or maintains MAD2L1, a MAD2 pseudogene, to kinetochores during spindle checkpoint activation induced by the spindle disruption [106]. Additionally, similarly to decreased mouse Nlrp5 mRNA expression with aging [57], there is also decreased mouse expression of Brca1 mRNA and protein in aged mouse oocytes [16], which could be a contributing factor underlying age-associated incidence of aneuploidy.

In our study, we observed reduced expression of BUB1, BUBR1, and TAp73 (which has been shown to modulate the activity of BUBR1) and increased retention of BRCA1, Aurora

89 kinase B, Aurora kinase C, and Dynactin p50 in Nlrp5-deficient oocytes suggesting that the Spindle Assembly Checkpoint may not be functioning adequately.

The spindle toxin, nocodazole, is commonly used to activate the SAC by preventing microtubule attachment to kinetochores. While nocodazole treatment caused the spindle to depolymerise, it also caused the destabilization of NALP5 protein from the chromosomes in MII oocytes. These results indicate that the correct attachment of the spindle to chromosomes is necessary to maintain localization of NALP5 on the chromosomal plate. Furthermore, this suggests that in ovulated oocytes, NALP5 may function as a sensing mechanism that wildtype oocytes elicit in response to detached spindles. Consequently, taken together with the NALP5 localization pattern, this data implies that when assessing the molecular pathway of the SAC in mature oocytes, NALP5 could be a crucial player that acts as a sensor of adequate kinetochore to microtubule attachment.

5.2 DNA DAMAGE In addition to the findings that we have obtained with the SAC analysis, there is also some evidence that Nlrp5 could be involved in DNA repair responses. The other means by which we investigated genome integrity in ovulated oocytes was by inducing DNA damage, via UV irradiation. To date, there is very limited information about the DNA damage/repair response in ovulated oocytes. Initially, we found that after UV exposure, wildtype oocytes displayed decreased NALP5 protein expression, cytoplasmically and specifically on the chromosomes. This pattern of expression was comparable to the NALP5 expression pattern noticed with the treatment of nocodazole, suggesting that both nocodazole and UV damage pathways appear to activate destabilization of NALP5.

In accordance with this, the high cytoplasmic staining of alpha-tubulin after UV irradiation, suggests that UV exposure causes partial depolymerisation of spindle microtubule filaments accompanied by increased frequency of oocytes with scattered chromosomes. This concurs with previous findings in human skin fibroblasts, in which it was concluded that UV damage to dimeric tubulin results in the disassembly of microtubules [146]. Taken together, these findings imply that UV exposure causes spindle damage in mature oocytes and also corroborates the results that increased BRCA1 expression after UV exposure was likely caused by spindle damage that was imposed by the UV irradiation.

We wanted to see if there were hallmarks of DNA damage in metaphase II-arrested oocytes and we additionally wanted to test whether Nlrp5-deficient oocytes also exhibited these hallmarks. We analysed this by performing the comet assay and by evaluating the protein accumulation of two DNA damage repair markers, 53BP1 and γH2AX, both of which are known to be associated with the sensing of DNA damage [124]. We observed that with UV damage, there was increased accumulation of 53BP1and γH2AX in MII oocytes. We further found that in Nlrp5-deficient oocytes, γH2AX and 53BP1 were elevated, indicative of DNA damage, although this was not detected by the comet assay. This raises the question of how sensitive the comet assay is in oocytes and whether it can detect nucleotide excision repair in these cells. If DNA damage is present in Nlrp5-deficient oocytes, the damage is below the threshold of detection by the comet assay. Because we saw evidence of UV irradiation causing spindle depolymerisation and chromosome scattering, we wanted to determine if the DNA damage markers could also be responding to the induced spindle damage. However, upon treatment of oocytes with nocodazole, causing spindle depolymerisation, we did not observe a change in the expression of 53BP1 nor γH2AX. This observation led us to rule out that the increased levels of 53BP1 and γH2AX are caused by spindle damage, but rather are likely caused by increased levels of DNA damage in the Nlrp5-deficient oocyte, despite our comet assay findings. Figure 27 highlights the UV damage pathway elicited in wildtype, ovulated oocytes.

UV Damage

Nucleotide/Base Spindle Damage/ Excision Repair Spindle Detachment

p ↑ H2AX ↑ 53BP1 ↓ NALP5 ↑ BRCA1

Figure 27: UV damage pathway in wildtype metaphase II-arrested oocytes. UV damage causes oocytes to elicit a form of excision repair, resulting in the increased accumulation of 53BP1 and γH2AX (phosphorylation of H2AX). BRCA1 increases after UV damage due to the secondary effects of increased spindle damage. NALP5 decreases after UV exposure as a result of increased spindle damage.

Recently, new exciting findings linking DNA repair with chromosomal decondensation in zygotes have been reported. Although, MII oocytes do exhibit DNA repair capabilities [113], however, the greatest evidence for repair occurring in gametes occurs in zygotes [35, 118, 135]. Present evidence suggests that in early zygotes there is equal DNA methylation in maternal and paternal genomes; however demethylation actively occurs in the paternal genome [30]. Up until now, DNA methylation has be thought of as a mark of epigenetic changes, however with these new findings we know that DNA methylation is likely present as a marker for DNA repair, as well [118]. More evidence indicates that the active mechanism of DNA repair happens at the zygote stage [118, 135], rather than in the oocyte stage. DNA demethylation could be thought of as a transient, intermediate step to DNA repair [118]. In mice, the paternal post-meiotic chromatin is believed to lack DNA repair capabilities [37], since the sperm only contributes the DNA to the zygote and not any DNA damage markers, such as: 53BP1, γH2AX, BRCA1 [30]. Subsequently, the paternal genome is repaired by the cytoplasmic machinery that is present in the oocyte. The factors produced by the oocyte are what repair that DNA. Consistently, there is likely some repair in female genome since eventually methylation also declines in this genome. Furthermore, in normal development, low levels of γH2AX is present, though, if an oocyte is fertilized with irradiated sperm, the foci size increases [37]. This validates the finding that it is the oocyte that is sensing and/or repairing the paternal genome, with the maternal factors present in its cytoplasm.

Condensation of chromatin, evidenced by phosphorylation of histone H3, is essential to ensure fidelity of chromosome segregation into daughter cells during cell division [73]. Interestingly, we found that phosphorylation of histone H3 (H3S10) increased at the 5 hours of culture timepoint compared to the 1 hour timepoint. This signifies that accumulation of phosphorylation on histone H3 is triggered in mouse oocytes by post-ovulatory aging and not by UV damage. Post-ovulatory aging in oocytes involves a gradual decrease of the physiological and biochemical processes that are essential for maintenance of developmental potential. This is accompanied by decreases in maturation promoting factor and mitogen-activated protein kinases [147]. Although, degradation of Nlrp5 oocyte transcripts was correlated with maternal age [57], our experiments showed that NALP5 protein did not appear to have an effect with post- ovulatory aging, since the 5 hour culture control group did not show a significant decrease in NALP5 expression compared to 1 hour controls. Our data suggest that the phosphorylation of histone H3 is altered by post-ovulatory aging in mouse oocytes.

Figure 28 shows a summary diagram displaying the changes of levels of various assayed- proteins in Nlrp5 deficiency at the metaphase II-arrested oocyte stage.

↓ Filia Nlrp5 KO

DNA Damage Spindle Damage ↓ Aurora A

↓ TAp73 ↑ Dynactin ↑ Aurora B ↑ Aurora C ↑ BRCA1 p50

p ↑ H2AX ↑ 53BP1 ↓ MAD2 ↓ BUB1 ↓ BUBR1

p ↑ H3 SAC

Meiosis progression without attached chromosomes

Developmental Female defect causing poor Multinucleation Infertility quality oocyte

Figure 28: Summary of the spindle and DNA damage-implicated roles of Nlrp5 in

Metaphase II-arrested oocytes.

6 FUTURE DIRECTIONS

Nlrp5 is a maternal-effect gene that is essential for the development of murine embryos. In this study, we have provided evidence that Nlrp5 has a role in maintaining genome stability in ovulated oocytes. Chromosomal instability in Nlrp5-deficient oocytes eventually resulted in the presence of micronuclei formation during the zygote stage of development. Micronuclei formation is a renowned indicator of extensive chromosomal damage [148]. One future direction would be to investigate the origin of the micronucleation defect since this would give insight into the mechanism of multinucleation or even aneuploidy [148]. It would be most informative to perform chromosomal spreads and investigate the karyotypes of Nlrp5-deficient ovulated oocytes. By especially looking at hyperploidy, we would be able to restrict chromosomal spread artifacts of preparation that may result in hypoploidy. This would allow us to assess aneuploidy in Nlrp5-deficient ovulated oocytes compared to wildtype oocytes and will allow us to decipher if multinucleation is eventually followed by aneuploidy in Nlrp5-deficient embryos. This technique will also allow us to use centromeres/kinetochore antibodies to get crisp localization of NALP5 and other markers in MII oocytes as many of these antibodies were not accessible to kinetochores, when whole mount immunocytochemistry was performed. Additionally, selected SAC- and DNA damage-involved target genes from our microarray analysis should be verified by real-time quantitative PCR analysis in wildtype and Nlrp5- deficient oocytes.

Another avenue that would be valuable to explore would be to assess whether the NALP5 protein physically interacts with any other SAC protein. As a result, it would be interesting to perform protein-protein interaction studies, by using an epitope-tagged protein of interest that would be permanently transfected in a cell line to conduct pull down experiments. Since we see localization of NALP5 to the kinetochores and because SAC proteins, BUB1 and BUBR1 also localize to this region of the chromosome, it would be interesting to see if NALP5 forms a complex with BUB1 or BUBR1. Moreover, it would also be useful to reconfirm the interaction between FILIA and NALP5, since in our study we see different localization patterns then previously described [47]. Furthermore, it would be informative to evaluate the functional role of Nlrp5 in the SAC, by using nocodazole at the metaphase I oocyte-stage, to assess whether Nlrp5 deficiency causes the premature chromosomal separation. This will allow us to see if

Nlrp5 deficiency abrogates the metaphase-arrest induced by nocodazole, indicating that NALP5 functions as a spindle checkpoint protein.

The comet assay is reliable in detecting major double stranded DNA damage; however this assay may not be the most suitable method for detecting UV-elicited DNA damage in MII oocytes. It has recently been discovered that mouse zygotes are capable of undergoing DNA repair by means of base excision repair [135]. In zygotes, this DNA repair has been linked to DNA demethylation changes and accumulation of γH2AX [118]. At the zygote stage, the pronuclear DNA decondenses and the chromatin becomes accessible, allowing for the investigation of DNA damage at this developmental stage by using assays, such as nick translation assays and BrdU and EdU incorporation assays. In contrast, in oocytes chromosomes are condensed and are less accessible; hence any enzyme-based assay is not a reliable indication of DNA damage. Additionally, it would be interesting to investigate DNA damage by measuring the accumulation of γH2AX and 53BP1 markers in Nlrp5-deficient zygotes, as this would give us conclusive evidence of whether DNA damage occurs with Nlrp5 deficiency. In addition, the extent of DNA methylation and whether there is a delay in the repair of paternal or maternal DNA could also be evaluated in Nlrp5-deficient zygotes, by performing immunocytochemistry.

There are newly emerging reports suggesting that increased aneuploidy is observed with increased maternal aging due to weakened centromere cohesion [149]. Since Nlrp5 decreases with aging [57], this could imply that Nlrp5 deficiency may lead to defective centromere cohesion. Furthermore, due to the fact that aging phenotypes are associated with aneuploidies and because Nlrp5-deficient mice are infertile, it would be valuable to look at the fecundity profile of heterozygote Nlrp5 mice by investigating litter sizes with aging.

Our findings thus far have focused on Nlrp5 phenotypes in mice. Ultimately, it would be useful to investigate whether Nlrp5 gene expression and NALP5 protein expression levels are altered in human aneuploid oocytes and embryos. Also, by collecting human multinucleated embryos from in vitro fertilization clinics, we could perform immunocytochemistry and evaluate whether there is differential NALP5 expression in these multinucleated embryos compared to control embryos. This could provide insight as to whether one of the major problems involved in human infertility, aneuploidy, is affected by Nlrp5.

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8 APPENDIX

Table 9: Complete list of mRNA transcripts with Decreased Expression in Nlrp5-Deficient MII Oocytes Gene Name Fold t-test Median SEM Median SEM change KO KO WT WT (p= ) (2 ^ Median Difference)

Nlrp5 NLR family, pyrin domain containing 5 8.21 0.00 7.64 0.20 10.67 0.39 Wfdc15a WAP four-disulfide core domain 15A 3.97 0.00 5.54 0.25 7.53 0.14 2610002 RIKEN cDNA 2610002J02 gene 3.58 0.00 3.84 0.15 5.68 0.14 J02Rik Rpl41 ribosomal protein L41 3.52 0.01 8.81 0.40 10.63 0.39 Zfp329 zinc finger protein 329 3.00 0.00 4.78 0.11 6.37 0.11 Rpl41 ribosomal protein L41 2.71 0.02 9.06 0.26 10.49 0.37 Nav2 neuron navigator 2 2.52 0.00 4.51 0.16 5.84 0.12 Epha5 Eph receptor A5 2.44 0.00 3.69 0.10 4.98 0.11 Tcf4 transcription factor 4 2.44 0.00 3.62 0.20 4.91 0.20 Zfp109 zinc finger protein 109 2.41 0.02 2.95 0.15 4.22 0.35 Basp1 brain abundant, membrane attached signal 2.38 0.00 6.85 0.13 8.10 0.10 protein 1 Lrrtm3 leucine rich repeat transmembrane neuronal 3 2.37 0.00 5.85 0.14 7.10 0.07 Xlr5c X-linked lymphocyte-regulated 5C 2.36 0.00 6.63 0.17 7.87 0.08 Cdkl2 cyclin-dependent kinase-like 2 (CDC2-related 2.34 0.05 5.22 0.38 6.45 0.24 kinase) Tcf4 transcription factor 4 2.32 0.00 5.42 0.17 6.63 0.14 Minpp1 multiple inositol polyphosphate histidine 2.31 0.00 7.01 0.22 8.22 0.13 phosphatase 1 Pggt1b protein geranylgeranyltransferase type I, beta 2.29 0.01 5.56 0.16 6.76 0.20 subunit Cftr cystic fibrosis transmembrane conductance 2.28 0.02 4.86 0.10 6.05 0.27 regulator homolog Flt1 FMS-like tyrosine kinase 1 2.27 0.00 7.02 0.13 8.20 0.14 Minpp1 multiple inositol polyphosphate histidine 2.24 0.00 5.06 0.15 6.22 0.05 phosphatase 1 Atg10 autophagy-related 10 (yeast) 2.22 0.00 5.67 0.16 6.83 0.08 Zfp420 zinc finger protein 420 2.22 0.04 3.00 0.13 4.16 0.42 Tcf4 transcription factor 4 2.22 0.02 4.50 0.23 5.65 0.39 Zkscan1 zinc finger with KRAB and SCAN domains 1 2.21 0.01 4.23 0.25 5.38 0.09 Zfp329 zinc finger protein 329 2.20 0.00 3.23 0.08 4.37 0.13

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Table 9: Complete list of mRNA transcripts with Decreased Expression in Nlrp5-Deficient MII Oocytes (Continued) Gene Name Fold t-test Median SEM Median SEM change KO KO WT WT (p= ) (2 ^ Median Difference)

Zkscan1 zinc finger with KRAB and SCAN domains 1 2.20 0.01 4.32 0.16 5.46 0.19 Plxna2 plexin A2 2.17 0.01 6.13 0.15 7.24 0.22 Rnaseh2 ribonuclease H2, subunit B 2.16 0.00 5.28 0.16 6.40 0.13 b Ocrl oculocerebrorenal syndrome of Lowe 2.14 0.01 6.61 0.11 7.71 0.19 4921506 RIKEN cDNA 4921506J03 gene 2.13 0.00 5.23 0.05 6.32 0.10 J03Rik Kif3b kinesin family member 3B 2.12 0.00 5.12 0.17 6.21 0.08 Ceacam carcinoembryonic antigen-related cell adhesion 2.12 0.00 9.64 0.21 10.72 0.10 20 molecule 20 Arpc5 actin related protein 2/3 complex, subunit 5 2.12 0.04 5.04 0.29 6.12 0.18 Zfp820 zinc finger protein 820 2.12 0.00 7.71 0.20 8.80 0.04 Mcl1 myeloid cell leukemia sequence 1 2.11 0.01 4.45 0.22 5.53 0.17 Pcyt1b phosphate cytidylyltransferase 1, choline, beta 2.11 0.03 6.22 0.29 7.29 0.08 isoform Slc5a3 solute carrier family 5 (inositol transporters), 2.07 0.01 3.18 0.18 4.23 0.23 member 3 4921506 RIKEN cDNA 4921506J03 gene 2.06 0.00 5.42 0.06 6.46 0.14 J03Rik Pde6b phosphodiesterase 6B, cGMP, rod receptor, beta 2.06 0.01 6.94 0.18 7.98 0.18 polypeptide 4933403 RIKEN cDNA 4933403F05 gene 2.05 0.01 5.30 0.15 6.34 0.17 F05Rik Zc3hav1 zinc finger CCCH type, antiviral 1 2.05 0.00 4.07 0.11 5.10 0.11 Eaf1 ELL associated factor 1 2.04 0.02 4.53 0.14 5.56 0.20 Leprotl1 leptin receptor overlapping transcript-like 1 2.04 0.02 4.72 0.23 5.74 0.30 Cd5 CD5 antigen 2.04 0.03 5.15 0.35 6.18 0.10 Zfp410 zinc finger protein 410 2.03 0.00 6.47 0.13 7.49 0.15 Mcl1 myeloid cell leukemia sequence 1 2.01 0.01 3.88 0.28 4.89 0.20 1810013 RIKEN cDNA 1810013L24 gene 2.01 0.00 9.56 0.06 10.57 0.04 L24Rik D3Ertd5 DNA segment, Chr 3, ERATO Doi 508, 2.00 0.00 9.28 0.07 10.28 0.04 08e expressed Fut11 fucosyltransferase 11 2.00 0.00 4.66 0.18 5.66 0.07

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Txnl1 thioredoxin-like 1 1.98 0.03 5.10 0.32 6.09 0.14 Pggt1b protein geranylgeranyltransferase type I, beta 1.98 0.00 7.21 0.15 8.20 0.05 subunit Xpr1 xenotropic and polytropic retrovirus receptor 1 1.97 0.00 4.48 0.10 5.46 0.06 Onecut1 one cut domain, family member 1 1.97 0.00 5.91 0.16 6.89 0.14 4921506 RIKEN cDNA 4921506J03 gene 1.97 0.00 6.08 0.12 7.05 0.05 J03Rik Eif2c3 eukaryotic translation initiation factor 2C, 3 1.97 0.01 5.42 0.20 6.39 0.06 Helz helicase with zinc finger domain 1.97 0.01 5.16 0.24 6.14 0.06 Nufip2 nuclear fragile X mental retardation protein 1.95 0.00 5.69 0.16 6.66 0.09 interacting protein 2 Gm6809 predicted gene 6809 1.95 0.02 3.83 0.11 4.79 0.34 Csnk1g1 casein kinase 1, gamma 1 1.95 0.02 5.80 0.34 6.77 0.14 Mtap9 microtubule-associated protein 9 1.93 0.02 7.48 0.19 8.43 0.22 Ptpn1 protein tyrosine phosphatase, non-receptor type 1.92 0.00 6.08 0.04 7.02 0.08 1 Tmem74 transmembrane protein 74 1.92 0.02 6.97 0.18 7.91 0.19 Ott ovary testis transcribed 1.91 0.02 4.99 0.25 5.93 0.17 Zfp330 zinc finger protein 330 1.91 0.01 8.92 0.23 9.85 0.11 Zfp248 zinc finger protein 248 1.91 0.04 3.15 0.26 4.08 0.21 Pura purine rich element binding protein A 1.91 0.02 3.77 0.13 4.70 0.24 Pja1 praja1, RING-H2 motif containing 1.90 0.05 3.31 0.22 4.24 0.31 Hspa13 heat shock protein 70 family, member 13 1.89 0.00 5.07 0.11 5.99 0.16 Bace1 beta-site APP cleaving enzyme 1 1.89 0.04 3.35 0.14 4.27 0.35 Mark4 MAP/microtubule affinity-regulating kinase 4 1.88 0.02 5.80 0.25 6.71 0.13 Tpm3 tropomyosin 3, gamma 1.87 0.01 8.71 0.22 9.61 0.12 B230354 RIKEN cDNA B230354K17 gene 1.86 0.00 4.07 0.13 4.97 0.14 K17Rik Ttn titin 1.86 0.01 4.00 0.18 4.90 0.14 Zkscan1 zinc finger with KRAB and SCAN domains 1 1.86 0.01 6.41 0.17 7.31 0.14 Tsga14 testis specific gene A14 1.86 0.01 7.04 0.15 7.93 0.27 Lcor ligand dependent nuclear receptor corepressor 1.84 0.01 8.96 0.13 9.84 0.17

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Gm1653 predicted gene 16536 1.84 0.00 4.96 0.13 5.84 0.07 6 Ide insulin degrading enzyme 1.84 0.01 5.53 0.21 6.41 0.04 Cbln1 cerebellin 1 precursor protein 1.84 0.01 3.66 0.24 4.54 0.14 4930539 RIKEN cDNA 4930539E08 gene 1.84 0.00 5.29 0.16 6.17 0.10 E08Rik 4933403 RIKEN cDNA 4933403F05 gene 1.84 0.01 5.03 0.19 5.90 0.14 F05Rik Raph1 Ras association (RalGDS/AF-6) and pleckstrin 1.83 0.00 6.38 0.15 7.26 0.11 homology domains 1 Tnrc6a trinucleotide repeat containing 6a 1.83 0.00 7.04 0.11 7.91 0.15 Ncdn neurochondrin 1.83 0.01 3.94 0.16 4.81 0.11 Dstyk dual serine/threonine and tyrosine protein kinase 1.82 0.00 8.80 0.10 9.66 0.15 Tnrc6a trinucleotide repeat containing 6a 1.82 0.00 8.33 0.13 9.20 0.06 Ppp1r7 protein phosphatase 1, regulatory (inhibitor) 1.82 0.00 5.45 0.06 6.31 0.08 subunit 7 Plxna2 plexin A2 1.82 0.00 6.96 0.10 7.82 0.13 Fam168 family with sequence similarity 168, member A 1.82 0.02 5.66 0.23 6.52 0.08 a St8sia6 ST8 alpha-N-acetyl-neuraminide alpha-2,8- 1.82 0.01 4.23 0.08 5.09 0.22 sialyltransferase 6 Pus7l pseudouridylate synthase 7 homolog (S. 1.81 0.00 7.43 0.16 8.29 cerevisiae)-like 2310047 RIKEN cDNA 2310047K21 gene 1.81 0.01 3.73 0.17 4.59 K21Rik Zfp263 zinc finger protein 263 1.81 0.00 3.79 0.09 4.64 Rnf168 ring finger protein 168 1.80 0.03 8.70 0.24 9.55 Fancl Fanconi anemia, complementation group L 1.80 0.03 3.59 0.15 4.44 Pdf peptide deformylase (mitochondrial) 1.79 0.00 3.10 0.11 3.94 Trps1 trichorhinophalangeal syndrome I (human) 1.79 0.01 5.68 0.17 6.52 Erc2 ELKS/RAB6-interacting/CAST family member 1.79 0.00 7.99 0.07 8.82 2 B3gnt2 UDP-GlcNAc:betaGal beta-1,3-N- 1.79 0.00 4.92 0.06 5.75 acetylglucosaminyltransferase 2 Zfp410 zinc finger protein 410 1.79 0.01 5.30 0.09 6.14

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Zfp108 zinc finger protein 108 1.79 0.02 4.69 0.22 5.53 Cstf2t cleavage stimulation factor, 3' pre-RNA subunit 1.79 0.03 5.89 0.26 6.73 2, tau Phf20 PHD finger protein 20 1.78 0.01 9.76 0.17 10.59 Chn2 chimerin (chimaerin) 2 1.78 0.02 5.85 0.21 6.68 2900009 RIKEN cDNA 2900009J20 gene 1.78 0.00 5.26 0.04 6.09 J20Rik Bicc1 bicaudal C homolog 1 (Drosophila) 1.77 0.00 6.67 0.06 7.49 Polrmt polymerase (RNA) mitochondrial (DNA 1.77 0.03 3.67 0.11 4.49 directed) Pglyrp1 peptidoglycan recognition protein 1 1.76 0.00 2.82 0.05 3.64 Aurkc aurora kinase C 1.76 0.00 11.87 0.08 12.68

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Table 10: Complete list of mRNA transcripts with Increased Expression in Nlrp5-Deficient MII Oocytes Gene Name Fold change t-test Median ST Median ST KO DEV WT DEV (2 ^ Median (p= ) KO WT Difference)

Hspa1b heat shock protein 1B 6.74 0.00 6.17 0.19 3.42 0.59 Hspa1b heat shock protein 1B 6.64 0.00 5.52 0.16 2.78 0.57 Hspa1b heat shock protein 1B 6.23 0.00 7.18 0.24 4.54 0.46 Tktl1 transketolase-like 1 5.84 0.00 6.20 0.51 3.65 0.21 Tktl1 transketolase-like 1 4.97 0.00 5.57 0.68 3.26 0.20 Hspb1 heat shock protein 1 4.44 0.00 10.56 0.32 8.41 0.18 Hspb1 heat shock protein 1 4.34 0.00 9.42 0.33 7.30 0.31 Stambpl STAM binding protein like 1 4.34 0.00 6.28 0.29 4.16 0.42 1 Rerg RAS-like, estrogen-regulated, growth-inhibitor 4.17 0.00 5.34 0.62 3.28 0.68 Dync1i2 dynein cytoplasmic 1 intermediate chain 2 3.87 0.00 10.04 0.27 8.09 0.68 Plac1 placental specific protein 1 3.65 0.00 7.20 0.48 5.34 0.60 Slc16a6 solute carrier family 16 (monocarboxylic acid 3.52 0.00 5.79 0.37 3.98 0.36 transporters), member 6 Hoxd1 homeobox D1 3.23 0.00 5.01 0.56 3.31 0.46 Spry4 sprouty homolog 4 (Drosophila) 3.17 0.00 8.64 0.22 6.97 0.44 H2afx H2A histone family, member X 3.14 0.01 8.52 0.36 6.87 0.63 Chic1 cysteine-rich hydrophobic domain 1 2.94 0.00 7.53 0.36 5.97 0.46 Pcdh15 protocadherin 15 2.93 0.00 6.83 0.37 5.28 0.35 Slc39a6 solute carrier family 39 (metal ion transporter), 2.90 0.01 9.53 0.43 7.99 0.60 member 6 Gm8947 predicted gene 8947 2.90 0.00 7.07 0.46 5.54 0.26 Slc35b4 solute carrier family 35, member B4 2.90 0.00 6.89 0.25 5.35 0.57 Sepp1 selenoprotein P, plasma, 1 2.87 0.01 5.87 0.69 4.35 0.42 Prl7d1 prolactin family 7, subfamily d, member 1 2.86 0.02 5.13 0.70 3.62 0.61 Kcnn2 potassium intermediate/small conductance 2.85 0.03 8.18 0.49 6.67 0.83 calcium-activated channel, subfamily N, member 2 Krt25 keratin 25 2.81 0.00 6.07 0.42 4.58 0.15 Fam32a family with sequence similarity 32, member A 2.81 0.00 7.19 0.27 5.70 0.50 Prr15 proline rich 15 2.76 0.00 5.51 0.30 4.04 0.19 Tmem15 transmembrane protein 159 2.74 0.00 5.96 0.14 4.50 0.50 9

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Table 10: Complete list of mRNA transcripts with Increased Expression in Nlrp5-Deficient MII Oocytes (Continued) Gene Name Fold change t-test Median ST Median ST KO DEV WT DEV (2 ^ Median (p= ) KO WT Difference)

B4galt5 UDP-Gal:betaGlcNAc beta 1,4- 2.73 0.04 8.05 0.27 6.60 0.83 galactosyltransferase, polypeptide 5 Abcd3 ATP-binding cassette, sub-family D (ALD), 2.71 0.04 5.03 1.07 3.58 0.18 member 3 Gli3 GLI-Kruppel family member GLI3 2.67 0.00 10.13 0.14 8.71 0.45 D13Ertd DNA segment, Chr 13, ERATO Doi 608, 2.63 0.00 4.92 0.43 3.53 0.26 608e expressed Cdc42 cell division cycle 42 homolog (S. cerevisiae) 2.58 0.00 5.52 0.27 4.15 0.33 Mup1 major urinary protein 1 2.55 0.00 5.15 0.33 3.79 0.15 Sesn3 sestrin 3 2.55 0.00 6.22 0.17 4.87 0.53 Pde12 phosphodiesterase 12 2.54 0.00 6.54 0.17 5.19 0.25 Bcor BCL6 interacting corepressor 2.52 0.01 5.33 0.20 4.00 0.52 Pcdh15 protocadherin 15 2.50 0.00 4.14 0.63 2.82 0.08 Hoxd8 homeobox D8 2.43 0.00 6.32 0.21 5.04 0.25 AF0670 cDNA sequence AF067063 2.43 0.02 4.95 0.60 3.67 0.51 63 Gli3 GLI-Kruppel family member GLI3 2.42 0.00 10.28 0.04 9.00 0.41 Ccne2 cyclin E2 2.41 0.02 6.64 0.80 5.37 0.35 Sbsn suprabasin 2.38 0.02 4.97 0.49 3.72 0.44 Mycbp c-myc binding protein 2.36 0.00 5.77 0.17 4.54 0.23 Cldn15 claudin 15 2.34 0.01 6.54 0.33 5.31 0.52 Pip5k1b phosphatidylinositol-4-phosphate 5-kinase, type 2.34 0.01 5.90 0.36 4.67 0.50 1 beta Rbmx2 RNA binding motif protein, X-linked 2 2.34 0.00 5.84 0.19 4.62 0.57 Scrn3 secernin 3 2.34 0.04 5.55 0.46 4.32 0.70 Hist3h2a histone cluster 3, H2a 2.31 0.01 5.07 0.52 3.86 0.32 Cebpd CCAAT/enhancer binding protein (C/EBP), 2.31 0.00 5.15 0.30 3.95 0.20 delta Cdh20 cadherin 20 2.28 0.02 5.66 0.37 4.48 0.74 Tlr3 toll-like receptor 3 2.26 0.00 5.02 0.21 3.84 0.10 Leo1 Leo1, Paf1/RNA polymerase II complex 2.26 0.02 6.91 0.48 5.73 0.47 component, homolog (S. cerevisiae) Glyr1 glyoxylate reductase 1 homolog (Arabidopsis) 2.25 0.02 8.05 0.17 6.88 0.56 Tfpi tissue factor pathway inhibitor 2.25 0.00 5.93 0.20 4.76 0.22

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6720489 RIKEN cDNA 6720489N17 gene 2.22 0.00 6.77 0.27 5.63 0.29 N17Rik Fam189 family with sequence similarity 189, member 2.20 0.00 6.57 0.33 5.44 0.26 a1 A1 Sec22a SEC22 vesicle trafficking protein homologe A 2.19 0.01 8.52 0.20 7.39 0.52 (S. cerevisiae) Ust uronyl-2-sulfotransferase 2.18 0.00 6.71 0.24 5.58 0.31 Pdha2 pyruvate dehydrogenase E1 alpha 2 2.17 0.03 6.62 0.59 5.50 0.58 2810422 RIKEN cDNA 2810422O20 gene 2.16 0.01 4.76 0.29 3.65 0.55 O20Rik Slc15a2 solute carrier family 15 (H+/peptide 2.14 0.00 8.35 0.31 7.25 0.30 transporter), member 2 Gm9898 predicted gene 9898 2.14 0.03 3.62 0.81 2.53 0.11 Prkcb protein kinase C, beta 2.13 0.00 4.63 0.26 3.54 0.29 Hsp90b1 heat shock protein 90, beta (Grp94), member 1 2.13 0.00 8.97 0.27 7.87 0.12 Cycs cytochrome c, somatic 2.13 0.00 8.03 0.11 6.94 0.31 Ank progressive ankylosis 2.12 0.01 8.97 0.10 7.89 0.65 Tmem33 transmembrane protein 33 2.12 0.01 6.90 0.33 5.81 0.60 Cd69 CD69 antigen 2.12 0.02 4.45 0.50 3.37 0.41 Il23a interleukin 23, alpha subunit p19 2.11 0.00 7.52 0.20 6.44 0.17 Cpox coproporphyrinogen oxidase 2.10 0.00 6.71 0.18 5.65 0.26 Tspan8 tetraspanin 8 2.09 0.04 5.27 0.66 4.20 0.40 Gbx2 gastrulation brain homeobox 2 2.09 0.03 6.14 0.32 5.08 0.56 Inpp5f inositol polyphosphate-5-phosphatase F 2.08 0.04 6.12 0.45 5.07 0.68 Snhg6 small nucleolar RNA host gene (non-protein 2.07 0.02 6.04 0.47 4.99 0.31 coding) 6 Ttc21b tetratricopeptide repeat domain 21B 2.07 0.00 5.61 0.30 4.56 0.37 Glul glutamate-ammonia ligase (glutamine 2.05 0.00 5.91 0.33 4.87 0.25 synthetase) Stk31 serine threonine kinase 31 2.05 0.00 7.83 0.39 6.80 0.20 Tlr3 toll-like receptor 3 2.04 0.00 7.21 0.14 6.18 0.30 Tpm4 tropomyosin 4 2.04 0.04 5.88 0.64 4.86 0.58 Eef2k eukaryotic elongation factor-2 kinase 2.02 0.00 9.09 0.32 8.07 0.17

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Cyp2g1 cytochrome P450, family 2, subfamily g, 2.02 0.01 5.12 0.50 4.11 0.13 polypeptide 1 Ehd4 EH-domain containing 4 2.02 0.01 7.72 0.18 6.71 0.50 Irx1 Iroquois related homeobox 1 (Drosophila) 2.02 0.02 4.96 0.35 3.95 0.40 Snapc3 small nuclear RNA activating complex, 2.02 0.02 5.91 0.15 4.90 0.70 polypeptide 3 Rybp RING1 and YY1 binding protein 2.01 0.02 7.80 0.48 6.79 0.40 Pnkd paroxysmal nonkinesiogenic dyskinesia 2.00 0.00 4.09 0.06 3.09 0.35 Doc2a double C2, alpha 2.00 0.02 4.21 0.45 3.21 0.49 Thoc1 THO complex 1 1.99 0.00 6.14 0.38 5.15 0.12 Clasp1 CLIP associating protein 1 1.98 0.01 5.77 0.43 4.78 0.32 A93001 RIKEN cDNA A930017M01 gene 1.98 0.05 5.98 0.73 4.99 0.49 7M01Ri k Plcb1 phospholipase C, beta 1 1.98 0.01 5.31 0.38 4.32 0.37 Stk31 serine threonine kinase 31 1.98 0.00 7.85 0.38 6.87 0.12 2500003 RIKEN cDNA 2500003M10 gene 1.98 0.01 6.28 0.19 5.30 0.42 M10Rik Dyrk1a dual-specificity tyrosine-(Y)-phosphorylation 1.98 0.01 6.81 0.21 5.83 0.47 regulated kinase 1a Siglec5 sialic acid binding Ig-like lectin 5 1.97 0.02 4.65 0.43 3.67 0.31 Cdc26 cell division cycle 26 1.97 0.02 9.32 0.31 8.34 0.43 Hist1h2a histone cluster 1, H2ae 1.97 0.03 8.36 0.66 7.38 0.34 e Gata6 GATA binding protein 6 1.97 0.01 5.49 0.37 4.52 0.52 Hhex hematopoietically expressed homeobox 1.96 0.03 8.10 0.60 7.12 0.28 Slc15a4 solute carrier family 15, member 4 1.96 0.02 6.78 0.14 5.81 0.55 Nedd4 neural precursor cell expressed, 1.96 0.00 6.05 0.21 5.08 0.35 developmentally down-regulated 4 Stard4 StAR-related lipid transfer (START) domain 1.94 0.02 5.21 0.14 4.25 0.51 containing 4 Pip5k1b phosphatidylinositol-4-phosphate 5-kinase, type 1.94 0.02 5.30 0.25 4.34 0.56 1 beta Edn1 endothelin 1 1.93 0.00 7.33 0.29 6.39 0.20 Mbp myelin basic protein 1.92 0.02 5.96 0.36 5.02 0.46

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Mrps6 mitochondrial ribosomal protein S6 1.92 0.01 4.73 0.34 3.79 0.34 Chrdl1 chordin-like 1 1.92 0.02 3.54 0.65 2.60 0.30 Ctla4 cytotoxic T-lymphocyte-associated protein 4 1.91 0.01 5.79 0.44 4.86 0.39 Nsun4 NOL1/NOP2/Sun domain family, member 4 1.91 0.01 9.95 0.19 9.02 0.40 Rnf167 ring finger protein 167 1.91 0.04 6.15 0.32 5.22 0.59 Pim2 proviral integration site 2 1.90 0.00 6.49 0.19 5.56 0.32 Rrh retinal pigment epithelium derived rhodopsin 1.90 0.01 6.13 0.37 5.20 0.42 homolog Foxc1 forkhead box C1 1.90 0.02 7.08 0.25 6.15 0.77 Pla2g12 phospholipase A2, group XIIB 1.90 0.00 6.59 0.31 5.66 0.08 b Dub1 deubiquitinating enzyme 1 1.90 0.04 3.61 0.62 2.68 0.22 Apoe apolipoprotein E 1.88 0.00 3.83 0.31 2.92 0.23 Mbp myelin basic protein 1.88 0.00 6.06 0.30 5.15 0.17 Mmadhc methylmalonic aciduria (cobalamin deficiency) 1.88 0.03 10.56 0.58 9.65 0.27 cblD type, with homocystinuria Hist1h3a histone cluster 1, H3a 1.87 0.00 7.01 0.32 6.11 0.20 Steap4 STEAP family member 4 1.87 0.03 8.22 0.44 7.32 0.56 Bdp1 B double prime 1, subunit of RNA polymerase 1.86 0.02 5.65 0.47 4.76 0.55 III transcription initiation factor IIIB Sfrs6 splicing factor, arginine/serine-rich 6 1.85 0.01 7.57 0.30 6.68 0.41 Mcm9 minichromosome maintenance complex 1.85 0.01 4.33 0.10 3.44 0.43 component 9 Pfn1 profilin 1 1.84 0.00 9.29 0.28 8.41 0.17 Eya4 eyes absent 4 homolog (Drosophila) 1.84 0.02 5.32 0.47 4.43 0.30 Spry4 sprouty homolog 4 (Drosophila) 1.84 0.01 8.30 0.44 7.42 0.18 Ttc33 tetratricopeptide repeat domain 33 1.84 0.04 7.26 0.55 6.37 0.34 Bank1 B-cell scaffold protein with ankyrin repeats 1 1.84 0.00 5.04 0.31 4.16 0.18 Ppp5c protein phosphatase 5, catalytic subunit 1.84 0.01 5.51 0.37 4.64 0.32 Fryl furry homolog-like (Drosophila) 1.84 0.02 7.10 0.18 6.22 0.51 Sohlh2 spermatogenesis and oogenesis specific basic 1.84 0.00 7.34 0.22 6.46 0.10 helix-loop-helix 2 Ubxn2b UBX domain protein 2B 1.83 0.02 4.82 0.19 3.94 0.64

126

Tfpi tissue factor pathway inhibitor 1.83 0.01 7.27 0.47 6.40 0.16 Actl6a actin-like 6A 1.83 0.01 6.99 0.27 6.12 0.33 Jam2 junction adhesion molecule 2 1.83 0.00 6.32 0.10 5.45 0.34 Fdx1 ferredoxin 1 1.82 0.01 10.75 0.27 9.88 0.26 Gm6792 predicted gene 6792 1.81 0.05 5.25 0.58 4.40 0.46 0610009 RIKEN cDNA 0610009D07 gene 1.81 0.01 8.19 0.30 7.33 0.32 D07Rik Ivns1abp influenza virus NS1A binding protein 1.81 0.02 7.02 0.28 6.17 0.52 Clcn2 chloride channel 2 1.79 0.02 4.43 0.34 3.58 0.50 Prim2 DNA primase, p58 subunit 1.79 0.01 8.95 0.29 8.10 0.31 Ms4a6d membrane-spanning 4-domains, subfamily A, 1.79 0.02 5.30 0.40 4.45 0.35 member 6D Mgea5 meningioma expressed antigen 5 1.79 0.00 5.11 0.11 4.27 0.31 (hyaluronidase) Rhobtb3 Rho-related BTB domain containing 3 1.79 0.02 7.75 0.35 6.91 0.39 Cpsf3 cleavage and polyadenylation specificity factor 1.79 0.00 7.20 0.10 6.36 0.30 3 Stim2 stromal interaction molecule 2 1.78 0.00 5.80 0.28 4.96 0.20 Ppp1r8 protein phosphatase 1, regulatory (inhibitor) 1.78 0.01 4.86 0.29 4.03 0.27 subunit 8 Mycbp c-myc binding protein 1.78 0.01 4.71 0.37 3.88 0.24 Lrrc28 leucine rich repeat containing 28 1.78 0.01 4.29 0.38 3.46 0.24 Ube4a ubiquitination factor E4A, UFD2 homolog (S. 1.78 0.04 6.27 0.18 5.44 0.49 cerevisiae) Cyp2f2 cytochrome P450, family 2, subfamily f, 1.78 0.00 4.95 0.12 4.12 0.24 polypeptide 2 Tcl1b4 T-cell leukemia/lymphoma 1B, 4 1.77 0.01 11.44 0.31 10.61 0.33 6720489 RIKEN cDNA 6720489N17 gene 1.77 0.02 6.65 0.26 5.83 0.36 N17Rik Slc35b4 solute carrier family 35, member B4 1.77 0.00 5.63 0.09 4.80 0.15 Ccl11 chemokine (C-C motif) ligand 11 1.77 0.01 5.41 0.20 4.59 0.38