Ministry of Higher Education And Scientific research University of Sulaimani Faculty of Medical Science School of Medicine Department of Anatomy

A Correlative Study of Oocytes Morphology with Fertilization, Cleavage, Embryo quality and Implantation rates After Intra Cytoplasmic Sperm Injection A thesis Submitted to the Council of the School of Medicine at the University of Sulaimani, in Partial fulfillment of the Requirements for the degree of MSc in embryology

By Tan Azad Salih M.B.Ch.B 2005

Supervised by Dr. Emad Ghanem Qassem M.B.Ch.B, MSc, F.I.C.M.S Assistant professor in Anatomy July 2014

Galawej 2714

Ramathan 1435

This thesis Is dedicated: To the memory of my beloved father…. it is your shining example that I try to emulate in all that I do. To Mom…. for her endless love, support, and encouragement To my sister….for her encouragement and expressed confidence in my abilities To Dana, Tina, Teni…….you are great love of my life

Acknowledgments I would like to show my gratitude to the University of Sulaimani for this opportunity……. I would like to sincerely thank my supervisor, Dr. Emad Ghanem Qassem for his guidance and support throughout this study. This study could not be completed without the assistance of Dr. Kani Muhammad Falah who has worked hard in the laboratory and provided guidance to me throughout this study. The present study would not have been accomplished without the help of Dr. Amanj Raheem Zangana and all the staff that they did working in Dwarozh-IVF center so I am grateful for all… Last, but never least, I must thank my unbelievably supportive husband Dana, our daughters (Tina, Teni) and my mother, who have demonstrated rare and amazing patience throughout my lengthy work and study….

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Abstract Background: Non-invasive selection of developmentally human oocytes may increase the overall efficiency of human assisted reproduction. Morphologic abnormalities in the oocyte are relevant for determining its developmental fate. Objective: to evaluate the influence of metaphase II (MII) oocyte morphology on intra cytoplasmic sperm injection (ICSI) outcomes. Design: A Correlative study. Setting: Dwarozh- IVF center. Patient(s): one hundred thirty two patients undergoing ICSI cycles and having female factors of infertility and unexplained infertility. Couples having male factors of infertility were excluded. Material and Methods: a total of 1200 oocytes were retrieved from 132 ICSI cycles, of which 1056 Metaphase II oocytes were evaluated. The criteria for morphological evaluations were: (i) Normal MII oocytes showing clear cytoplasm with uniform texture and homogenous fine granularity, a round or ovoid first polar body with a smooth surface, and perivitelline space of normal size. (ii) MII oocytes with extra cytoplasmic abnormalities (first polar body and perivitelline space abnormalities). (iii) MII oocytes with cytoplasmic abnormalities (dark cytoplasm, granular cytoplasm, inclusion body and presents of vacuoles). (iv) MII oocytes with combined cytoplasmic and extra cytoplasmic abnormalities. Oocytes with shape abnormalities were not recorded. Embryos transferred day 2 or 3 after injection.

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Main Outcome Measure(s): Fertilization, Cleavage, Embryo quality, and Implantation rates. Result(s): from 1056 MII oocytes, 180 (17.04%) had normal morphology while 876 (82.95%) had at least one demonstrable morphological abnormality. Cytoplasmic abnormalities (dark cytoplasm, granular cytoplasm, presence of inclusion body and vacuoles) were observed in 516 (58.9%) of the oocytes. Extra cytoplasmic abnormalities (anomalies of polar body and perivitelline spaces) were observed in 104 (11.87%) while combined cytoplasmic and extra cytoplasmic abnormalities were responsible for the remaining 256 (29.22%). Mean age of patients were (33.98±5.581). There were no significant differences in fertilization, cleavage, and embryo quality between the groups but there was highly significant difference in implantation rate between them. Conclusion: MII oocyte morphology did not affect fertilization, cleavage, and embryo quality including high quality embryo, but implantation rate was higher in the group with normal oocyte morphology than the group of abnormal oocyte morphology.

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Table of Contents Chapter One 1. Introduction............................................................................................. 1 1.1 The development of the oocyte ......................................................... 6 1.1.1 Primordial germ cells: the origin of oocytes .............................. 6 1.1.2 Transition of oogonia to primary oocytes................................... 8 1.1.3 Formation of primordial follicles ............................................... 9 1.1.4 Transition of primordial follicles into primary follicles (initiation of follicle growth) ............................................................. 10 1.1.5 Follicle growth to pre-antral and antral stages ......................... 12 1.2 The oocyte ....................................................................................... 14 1.2.1 Cumulus-enclosed oocytes ....................................................... 16 1.2.2 Oocyte maturation stage ........................................................... 18 1.2.3 Oocyte size and shape ............................................................... 20 1.2.4 Cytoplasmic features ................................................................ 21 1.2.5 Extra cytoplasmic features ........................................................ 23 a. zona pellucida………………………….…………………..23 b. perivitelline space………………………………………….25 c. polar body……………………...…………………………..26 1.3 Ovulation ......................................................................................... 28 1.4 Fertilization ..................................................................................... 31 1.5 The cleavage stage embryo ............................................................. 35 1.5.1 Cell numbers ............................................................................. 36 1.5.2 Fragmentation ........................................................................... 37 IV

1.5.3 Blastomere size ......................................................................... 38 1.5.4 Nucleation ................................................................................. 39 1.6 Implantation .................................................................................... 41 1.6.1 The proliferative phase ............................................................. 42 1.6.2 The secretory phase .................................................................. 43 1.6.3 Endocrinological aspects .......................................................... 43 1.6.4 Apposition, adhesion and penetration ...................................... 44 Chapter Two 2. Patients and methods ............................................................................ 46 2.1 Ethical consideration ....................................................................... 46 2.2 Study design .................................................................................... 46 2.3 Ovarian stimulation ......................................................................... 47 2.4 Oocyte retrieval ............................................................................... 48 2.5 Oocyte denudation........................................................................... 48 2.6 Oocyte evaluation............................................................................ 49 2.7 Semen preparation ........................................................................... 51 2.8 Intra cytoplasmic sperm injection procedure .................................. 52 2.9 Assessment of fertilization, embryo cleavage and embryo transfer ............................................................................................................... 53 2.10 statistical analysis .......................................................................... 54 Chapter Three 3. Results................................................................................................... 57 3.1 General characteristics: ................................................................... 57

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3.2 Influence of MII Oocytes Morphology on Fertilization, cleavage and embryo quality rates ....................................................................... 60 3.3 Comparisons between normal MII oocytes and MII oocytes with cytoplasmic abnormalities in relation to fertilization, cleavage and embryo quality....................................................................................... 61 3.4 Comparisons between Normal MII oocytes and MII oocytes with extra cytoplasmic abnormalities in relation to fertilization, cleavage and embryo quality....................................................................................... 62 3.5 Comparison between Normal MII oocytes and MII oocytes with combined abnormalities in relation to fertilization, cleavage and embryo quality....................................................................................... 63 3.6 Relationship between MII oocytes morphology and implantation rate ......................................................................................................... 64 Chapter Four 4.1 Discussion ....................................................................................... 74 4.2 Conclusion:...................................................................................... 79 4.3 Recommendations: .......................................................................... 80 References................................................................................................. 81 Appendix………………………………………………………………… ……………………………………………. Questionnaire Ethic committee approval Arabic abstract Kurdish abstract

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List of Tables Table (3.1): General Characrestics of ICSI cycles ................................... 58 Table (3.2): Distribution of abnormal MII oocytes according to their morphology ............................................................................................... 59 Table (3.3): Fertilization, Cleavage and Embryo quality rates after injection of normal and morphologically abnormal MII oocytes ............ 60 Table (3.4): Compared results between normal MII oocytes morphology and MII oocytes with cytoplasmic abnormalities after injection ............. 61 Table (3.5): Compared results between normal MII oocytes morphology and MII oocytes with extra cytoplasmic abnormalities after injection .... 62 Table (3.6): Compared results between normal MII oocytes and MII oocytes with combined abnormalities after injection ............................... 63 List of Figures Figure (3.1): Copmarison of implantation rate between normal MII oocytes morphology and abnormal MII oocyte morphology ................... 64 Figure (3.2): Comparison of implantation rate between normal MII oocytes morphology and MII oocytes with cytoplasmic abnormalities ... 65 Figure (3.3): Comparison of implantation rate between normal MII oocytes and MII oocytes with extra cytoplasmic abnormalities .............. 66 Figure (3.4): Comparison of implantation rate between normal MII oocytes morphology and MII oocytes with combined abnormalities ...... 67 List of Plates Plate (2.1) Double Lumen Aspiration Needle .......................................... 55 Plate (2.2) Nikon Stereo Microscope ....................................................... 55 Plate (2.3) Inverted ICSI Microscope ....................................................... 56 Plate (2.4) CO2 incubator ......................................................................... 56

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List of Micrographs Micrograph (3.1) Immature oocyte at Germinal vesicle stage ………..68 Micrograph (3.2) Immature oocyte at Metaphase I stage …….……….68 Micrograph (3.3) Normal mature oocyte at Metaphase II stage……….68 Micrograph (3.4) Abnormal MII Oocytes with cytoplasmic inclusion body…………………………………………………………………….69 Micrograph (3.5) Abnormal MII Oocytes with cytoplasmic vacuoles…69 Micrograph (3.6) Abnormal MII oocytes with central granulation of cytoplasm………………………………………………………………69 Micrograph (3.7) Abnormal MII oocytes with granular cytoplasm…....70 Micrograph (3.8) Abnormal MII oocyte with large and small polar body…………………………………………………………………….70 Micrograph (3.9) Abnormal MII oocyte with fragmented polar body and central granulation…………………………………………………..…70 Micrograph (3.10) Abnormal MII oocyte with dark zona pellucida and large PVS…………………………………………..…………………..71 Micrograph (3.11) Abnormal MII oocyte with PVS debris………..….71 Micrograph (3.12) Abnormal MII oocyte with large polar body and large PVS……………………………………………………………………..71 Micrograph (3.13) Abnormal MII oocyte with dark cytoplasm and PVS debris……………………………………………………………………72 Micrograph (3.14) Zygote obtained by ICSI of oocyte displaying 2 pronuclei………………………………………………………………..72 Micrograph (3.15) Day 2 high quality embryo………………………….73 Micrograph (3.16) Day 2 low quality embryo…………………………..73 Micrograph (3.17) Day 3 high quality embryo…………………….……73 Micrograph (3.18) Day 3 low quality embryo…………………………..73 VIII

List of abbreviations AMH………………………………………….…Anti mullerian hormone ART……………………………………..Assisted reproductive technique CC……….………………………………………………..…...Corona cell COC…….………………………………...…….Cumulus-oocyte complex E2…………………………………………….…….………..…Estradiol 2 ECM………………………………………...……….Extra cellular matrix FSH……………………………………….Follicular stimulating hormone GC……...……………………………………………..……Granulosa cell GnRH……………………………….…..Gonadotropin releasing hormone GnRHa…………………………Gonadotropin releasing hormone agonist GV……………………………………………….……....Germinal vesicle GVBD…………………….……………..….Germinal vesicle break down HCG………………………….…….……Human chorionic gonadotrophin ICSI………………………………….…Intra cytoplasmic sperm injection IVF………………………….…………………….…..In-vitro fertilization LH…………………………….……………………..Luteinizing hormone LIF……………………………..…….………..Leukemia-inhibitory factor MI…………………………………………………………….Metaphase I MII…………………………………………..……………….Metaphase II IX

MS………………………………….….……………..…..Meiotic spindle NPBs……………………………….……...……Nuclear precursor bodies OHSS………………………………Ovarian hyper stimulation syndrome P4………………………………..…….………….…………Progesterone PBI……………………………………………...…….……...Polar body I PGC…………………………………………………Primordial germ cell PGs……………………………………….…...….…………Prostaglandin PN……………………………………………….…….………Pronucleus PVS…………………………………………….………Perivitelline space SER………………………………...…..…Smooth endoplasmic reticulum TGF-b…………………………….……..Transforming growth factor-beta ZP………………………………….………….……………Zona pellucida ZPTV…………………………….……Zona pellucida thickness variation

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Chapter one Introduction and literature review

1. Introduction Infertility is defined as the inability of a couple to achieve conception or bring a pregnancy to term after 12 months or more of regular unprotected sexual intercourse (WHO, 2010). It is a growing concern affecting up to 15% of couples trying to conceive globally (Leisegang et al., 2014). The prevalence varies widely, being less in developed countries and more in developing countries where limited resources for investigation and treatment are available (Kamel, 2010). According to literature survey, the most common causes of infertility are: (1) Male factor (Irvine, 2004; Olatunji and Sule-odu, 2003; Bayasgalan et al., 2004). (2) Female factors such as ovulation dysfunction, scarring from inflammatory or infectious diseases, nutrient deficiencies, hormone imbalance, ovarian cysts, and transport system abnormality from cervix through fallopian tube (Devoto et al., 2009; Nichi et al., 2011). (3) Combined female and male factor, and unexplained infertility (Poppe and Velkeniers, 2002; Ikechebula et al., 2003; Bayasgalan et al., 2004). With the fast progression in reproductive medicine and the experiences gained through infertility management, a wider range of treatment options have become available to infertile couples (Jose-Miller et al., 2007; Oladokun et al., 2009;Macaluso et al., 2010). There are three main types of infertility treatment: medical treatment (such as ovulation induction therapy); surgical treatment (such as laparoscopy and hysteroscopy); and the different assisted reproduction techniques (Brinsden et al., 1998). Choices of infertility treatment are often related to issues of efficacy, cost, ease of use or administration, and its side effects. Legal, cultural and religious inquiries have limited the available choices in some countries, 1

such as the use of donor sperms or oocytes. Treatment options available for any particular infertile couples also depend on the duration of infertility, the affected partner, the age of the female partner and if any of the partners have a previous child, in addition to the underlying pathological cause (Kamel, 2010). Assisted reproductive technologies (ART) have been developed in order to overcome both female and male infertility. Intra cytoplasmic sperm injection (ICSI) had been widely used to treat couples with infertility because of severely impaired sperm characteristics and for whom in-vitro fertilization (IVF) had failed (Xia, 1997). The cause of fertilization failure in IVF is most often due to difficulty of sperm penetration (Dyban et al., 1992), but the reason why fertilization does not occur when intra cytoplasmic sperm injection (ICSI) is applied is not clear. Lack of fertilization with ICSI may be attributed to the deficiency of spermassociated oocyte activating factor (Dozortsev et al., 1995). However, some intrinsic oocyte problems may also be responsible for fertilization failure. Oocytes abnormalities that can be assessed at the light microscopy level may be associated with failure of fertilization in the presence of apparently normal spermatozoa (Balaban and Urman, 2006). In assisted reproductive treatment programs, controlled ovarian hyper stimulation for multiple follicular developments has become widely used this allows the recovery of numerous pre ovulatory oocytes for ART treatment program (Pelletier et al., 2004). Little attention has been focused on oocyte morphology in standard IVF techniques, because it is often difficult to assess the cytoplasmic morphology of the oocyte and the exact stage of maturation as the

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oocytes are always surrounded by cumulus oopheros or corona cells at the time of collection (Serhal et al., 1997). Following the removal of the cumulus–corona cells in preparation for intra cytoplasmic sperm injection (ICSI), oocyte evaluation is more accurate. Oocyte maturation is based on the nuclear maturation status, the morphology of the cytoplasm and on the appearance of the extra cytoplasmic structures. The presence of the first polar body (IPB) is normally considered to be a marker of oocyte nuclear maturity. However, recent studies using polarized light microscopy have shown that oocytes displaying a polar body may still be immature (Rienzi et al., 2005). Only those displaying a meiotic spindle (MS) can in fact be considered as true, mature, Metaphase II (MII) stage oocytes. Nuclear maturity alone is, in fact, not enough to determine the quality of an oocyte. Nuclear and cytoplasmic maturation should be completed in a coordinated manner to ensure optimal conditions for subsequent fertilization (Rienzi et al, .2012). An ideal mature human oocyte, based on morphological characteristics, should have a ‘normal-looking’ cytoplasm, a single polar body, an appropriate zona pellucida (ZP) thickness and proper perivitelline space ( Swain and Pool, 2008). However, the majority of the oocytes retrieved after ovarian hyper stimulation exhibit one or more variations in the described ‘ideal’ morphological criteria (Ebner et al., 2006; Rienzi et al., 2008). This is also true for oocytes obtained from proven fertile donors (Ten et al., 2007). Oocytes quality and its influence on fertilization rate and embryo development is still a matter of controversy in ART cycles (Khalili et al., 2005). Kahraman et al., (2000) reported oocyte morphology to be an 3

important prognostic factor in successful treatment of ICSI cases. Alikani et al., (1995) also reported a higher rate of miscarriage in women with dysmorphic oocytes. On the other hand, De Sutter et al., (1996) reported that oocyte morphology does not correlate with fertilization rate and embryo quality after ICSI. To date, the extent to which the morphology of the oocyte at the light microscopy level correlates with the results of ICSI is controversial.

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Aim of the study: The main purpose of this study is to detect the influence of MII oocytes morphology on:  Rates of fertilization, cleavage, and embryo quality.  Implantation rate.

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1.1 The development of the oocyte Ovarian reserve is determined by the number of primordial follicles in the ovary. Quiescent primordial follicles are activated for growth and pass through stages of development before they reach the antral stage. Then a cohort of antral follicles is recruited for further growth, dominance and ovulation under the cyclic stimulation of gonadotrophins. What triggers the initiation of growth in primordial follicles has remained a mystery for decades (Oktem and Urman, 2010). 1.1.1 Primordial germ cells: the origin of oocytes The earliest primordial forms of germ cells (primordial germ cells, PGCs) are differentiated from the proximal epiblast adjacent to the extraembryonic ectoderm (Ying et al., 2001). The onset of germ cell competence is marked by the expression of an interferon inducible transmembrane protein (fragilis) on the germ cells. Fragilis subsequently induces the expression of Stella, a gene expressed exclusively in Lineage-restricted germ cells, allowing escape from somatic cell fate and retention of pluripotency as they migrate through the hindgut mesentery and arrive in the gonad (Lange et al., 2003). Primordial germ cells (PGCs) first appear as a cluster of approximately 100 cells in the endoderm of the dorsal wall of the yolk sac near the allantois between the third and fourth weeks of gestation in the human then Primordial germ cells (PGCs) migrate to the hindgut and dorsal mesentery during the fourth and fifth weeks of gestation, respectively (Mc et al., 1953).

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By the seventh week of gestation, colonization of gonadal tissue by germ cells is complete. Germ cells are essential for the formation and maintenance of the ovary: in their absence the gonad degenerates into cordlike structures (Merchant-Larios and Centeno, 1981). Once the Primordial germ cells (PGC) have arrived in the gonad, they undergo more extensive proliferation such that their number rapidly increases from merely 10, 000 at the sixth week of gestation to 600,000 at the eighth week. With rapid mitotic activity, their number further rises to 6 million at the 20th week of gestation. Thereafter the rate of oogonial mitosis progressively declines and ends at 28 weeks with an almost equally increasing rate of oogonial atresia, which peaks at 20 weeks of gestation (Oktem and Oktay, 2008). During childhood most oocytes become atretic only approximately 400, 000 are present by the beginning of puberty and fewer than 500 will be ovulated (Sadler, 2006).

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1.1.2 Transition of oogonia to primary oocytes Primordial germ cells (PGCs) are called oogonia once they reach the gonads. The oogonia exhibit higher mitotic activity compared with PGCs and undergo several rounds of mitotic divisions prior to meiosis. Mitotic activity of the oogonia is the major determinant of the oocyte pool (Oktem and Urman, 2010). The last several rounds of mitosis before the initiation of meiosis gives rise to the formation of a syncytium (clusters of oogonia) in which oogonia are connected to each other by their cell membranes, the cytoplasmic bridges is caused by incomplete cytokinesis during cell division. In oogonia connected to each other with cell bridges, meiosis begins simultaneously, suggesting propagation of the signal triggering meiotic entry through these bridges. Initiation of pre meiotic DNA synthesis marks the end of the oogonial stage; thereafter germ cells are called primary oocytes. Entry into meiosis begins between 8 and 13 weeks of gestation well before follicle formation (Oktem and Urman, 2010). All surviving primary oocytes enter prophase of meiosis I, and most of them are individually surrounded by a layer of flat epithelial cells (Sadler, 2006).

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1.1.3 Formation of primordial follicles The first primordial follicles appear in the human fetus as early as 15th week of gestation and are complete by 6 months after birth (McGee and Hsueh, 2000; Maheshwari and Fowler, 2008). Primordial follicles are composed of diplotene oocytes (30–60 µm) surrounded by flattened pre granulosa cells. The dictyate is a protracted stage of arrest, in which the primordial follicles remain in prophase of MI and remain there until meiosis is, resumed (Fragouli et al., 2014). Transcriptomic studies in human and rodents have identified a variety of genes involved in primordial follicle assembly, such as transcription factors zona proteins, meiosis-specific enzymes and nerve growth factors. Initiation of meiosis in the oogonia (becoming oocytes) with investment of granulosa cells to form the primordial follicle appears to provide protection from atresia, as the oogonia cannot persist beyond the seventh month of gestation without entering meiosis. Therefore, ovaries in newborn are usually devoid of oogonia (Abir et al., 2002).The reproductive life span of women is determined by the number of primordial follicles in the ovary. At present there is no hormonal or any other marker of primordial follicles to be used clinically for the prediction of ovarian reserve (Oktem and Urman, 2010).

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1.1.4 Transition of primordial follicles into primary follicles (initiation of follicle growth) Primordial follicles remain in a dormant phase until being recruited into the primary stage for growth. This takes place with the onset of puberty in females and the initiation of the menstrual cycle (reviewed in Huang et al., 2012). Traditional thinking proposes that rather than a single signaling pathway, an orchestrated multi-directional communication among the oocytes and somatic cells (granulosa cells and thecal cells), and certain extra-cellular matrix components and growth factors acting in an autocrine and paracrine manner, play roles in this transition and subsequent growth of follicles (Eppig, 2001; Skinner, 2005). Anti-Mullerian hormone (AMH, also known as Mullerian inhibiting substance) is a member of the transforming growth factor-beta (TGF-b) family and is released from granulosa cells of growing follicles. There is an increased recruitment of primordial follicles into the growing pool in AMH null mice, suggesting a negative effect of AMH on the primordialto-primary follicle transition (Durlinger et al., 1999; Durlinger et al., 2002). AMH is produced by the granulosa cells of growing pre-antral and small antral follicles as a dimeric glycoprotein (Visser and Themmen, 2005). AMH has the least inter- and intra-cycle variability, thus making it a good marker for evaluation in random blood samples; it also correlates well with the number of antral follicles in the ovary and the number of oocytes retrieved (Van Rooij et al., 2002; Ebner et al., 2006).

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Flattened granulosa cells of primordial follicles become cuboidal during transition into the primary stage along with an increase in oocyte diameter (>60 µm) and the acquisition of zona pellucid (Rankin et al., 1996).The initial recruitment of quiescent primordial follicles into the growing pool as primary follicles starts in fetal life and continues postnatally until the ovarian reserve is depleted, leaving behind only ~1000 primordial follicles in the ovary at the time of menopause (Oktem and Oktay, 2008). The process is continuous and different from the cyclic recruitment of a cohort of antral follicles under the actions of FSH. In addition to inhibitory signals that inhibit premature activation of primordial follicles, there are some other signals in the ovary that promote the transition of primordial follicles to primary follicles. With coordinated and synergistic actions of these signals arising from different compartments, such as oocytes, somatic cells and stroma, growth is initiated in primordials. This could also explain why isolated primordial follicles do not survive in culture, but grow in situ in ovarian tissue culture (O’Brien et al., 2003). Furthermore, FSH is not required for this transition as primordial follicles do not express FSH receptors (Oktay et al., 1997).

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1.1.5 Follicle growth to pre-antral and antral stages As the primary oocyte begins to grow, surrounding follicular cells change from flat to cuboidal and proliferate to produce a stratified epithelium of granulosa cells, and the unit is called a primary follicle. Granulosa cells rest on a basement membrane separating them from surrounding stromal cells that form the theca folliculi. Moreover, granulosa cells and the oocyte secrete a layer of glycoproteins on the surface of the oocyte, forming the zona pellucida (Sadler, 2006). An increase in oocyte diameter and formation of basal lamina, zona pellucida and theca cell layer are among the other changes that characterize this developmental stage (Knight and Glister, 2006). During this phase, follicle diameter increases from 40–60 µm at the primary stage and to 120–150 µm at the pre-antral stage. With further growth, follicles reaching a diameter of 200 µm enter the antral stage. It is also during this stage that the follicle begins to exhibit some fluid-filled spaces within the granulosa cell layers, which coalesce to form the antral cavity, along with increased vascularization of the theca layer, continued growth of oocytes and proliferation of granulosa and theca cells. Development of a multi-layered secondary follicle from a primary follicle with a single layer of granulosa cells is a long process that takes months in humans. This slow process appears not to be mediated by the actions of gonadotrophins even though pre-antral follicles may express FSH receptors (Oktay et al., 1997).

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Under the influence of gonadotrophins [follicle stimulating hormone (FSH) and luteinizing hormone (LH)] MI is resumed; the oocyte progresses to metaphase I; chiasmata are resolved and homologous chromosomes separate; one set (n = 23) remains in the oocyte (secondary oocyte) while the other enters the first polar body (PB) (reviewed in Sutton et al., 2003; Fragouli et al., 2011). The oocyte continues on, entering MII before arresting again at the second metaphase. MII is not completed until the oocyte is fertilized (Sadler, 2006). Granulosa cells have no direct blood supply. The basal lamina works as a blood-follicle barrier and separates granulosa cells from the vascularized thecal cell layer necessitating an intimate contact between neighboring granulosa cells and the oocytes. An extensive network of gap junctions between granulosa cells couples them into an integrated functional syncytium. These junctions are composed of connexin proteins arranged in a hexameric configuration and allow not only an effective communication but also efficient metabolic exchange and transportation of molecules between granulosa cells. Granulosa cells also communicate with the oocytes via the gap junctions projecting through the zona pellucida to the plasma membrane of the Oocytes (Oktem and Urman, 2010).

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1.2 The oocyte The female gamete plays a crucial role in determining embryo competence and therefore in vitro fertilization (IVF) results. Oocyte quality is not only influenced by the nuclear and mitochondrial genome, but also by the microenvironment provided by the ovary and the preovulatory follicle that influences transcription and translation, and as a consequence, cytoplasmic maturity (Rienzi et al., 2012). The acquisition of oocyte competence relies on a complex cascade of events that occurs during follicular development. In the course of acquiring these competencies, cytoplasmic changes occur that include mRNA transcription, protein translation, posttranslational modification of proteins and ultra-structural changes (Sirard et al., 2006). Successful completion of these events is independent of nuclear maturation and is collectively referred to as cytoplasmic maturation (Krisher, 2004). As an oocyte grows and matures, it acquires the ability to resume and complete meiosis (First et al., 1988), successfully undergo the fertilization process, initiate and sustain embryonic development (Sirard et al., 2006). An oocyte that has not completed cytoplasmic maturation is of poor quality, and thus unable to successfully complete normal developmental processes. However, the mechanisms that impair oocyte developmental competence are still unclear (Krisher, 2004). Application of ovarian stimulation in human reproduction further complicates the situation. In contrast to the in vivo process, where oocyte maturation occurs as the result of a natural selection procedure (Rienzi et al., 2008), in stimulated cycles, pharmacologic doses of gonadotrophins create a supraphysiologic hormonal environment that induces the growth 14

of a cohort of follicles, which, under natural conditions, would become atretic and regress (Figueira et al., 2010) allowing successful maturation of oocytes with compromised quality (Swain and Pool 2008). Nuclear maturity alone is neither sufficient to determine the competence of an oocyte, nor to ensure that the meiotic apparatus of the oocyte has progressed correctly to metaphase II (MII) stage (De Santis et al., 2005). A deficiency in cytoplasmic maturation could compromise all processes that prepare the oocyte for activation and adequate fertilization. Moreover, the morphological appearance of the oocyte may indicate the developmental potential of the subsequent embryo (Rienzi et al., 2008). With the advent of intra cytoplasmic sperm injection (ICSI), which requires the denuding of the oocyte from the cumulus corona cells, a wide range of information regarding oocyte morphology has emerged and permits the oocyte to be observed immediately before sperm injection using an inverted microscope. It is generally accepted that good-quality human MII oocytes should have a clear, moderately granular cytoplasm that does not contain inclusions, a small perivitelline space (PVS) containing a single unfragmented first polar body and a round, clear, colorless zona pellucida (ZP) (Van Blerkom, 1990;Rienzi et al., 2008). Nevertheless, more than half of all collected oocytes show at least one morphological abnormality (De Sutter et al., 1996; Ebner et al, .2006). Morphological abnormalities of oocytes are most commonly classified as either intra cytoplasmic features (increased cytoplasmic granularity and presence of cytoplasmic inclusions) or extra cytoplasmic features (large perivitelline space (PVS), PVS granularity, and fragmented or irregular first polar body). (Figueira et al., 2010).

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1.2.1 Cumulus-enclosed oocytes During follicular antrum formation, granulosa cells (GCs) differentiate into mural GCs lining the follicular wall, and corona cells surrounding the oocyte. Within the cumulus mass, corona cells in close contact with the oocyte develop cytoplasmic projections which cross the zona pellucida and form gap junctions with the oolemma. This organized structure is called the cumulus–oocyte complex (Albertini et al., 2001). In natural spontaneous cycles, oocyte nuclear maturation runs parallel to the gradual FSH-dependent expansion of the cumulus and corona cells, whereas this synchrony may be disturbed in stimulated cycles (Laufer et al., 1984). The oocyte is known to depend on its surrounding somatic cells for a myriad of activities ranging from sources of signaling molecules to metabolites (Hussein et al., 2006; Su YQ et al., 2008). Interestingly, the interdependence between the two cell compartments of the cumulus– oocyte complex (COC) is reciprocal. Indeed, the cumulus cells also depend on the oocyte for their normal differentiation, regulation, and functions (Gilchrist et al., 2004; Gilchrist et al., 2008). Mechanistically, the development of cumulus cells and oocyte appear coordinated through a complex and regulated set of intercellular interactions, including direct cell–cell contacts, gap junctional communications, and paracrine signaling (Gilchrist et al., 2008). Specialized cellular projections (from the cumulus cells to the oocyte surface) permit physical and close link with the oocyte. These transzonal projections exist in cumulus oocyte complex( COCs) of all mammalian systems described to date; they are rich in either microtubules or microfilaments, and are reminiscent of cell processes in neurons, themselves examples of another cell type that rely extensively on cell communication and support from other cells. Gap 16

junctions exist between cumulus cells and oocytes, and their functional roles are supported by genetic programming and several in vitro lines of evidence (Kidder and Mhawi, 2002). Another essential component of the cumulus oocyte complex (COC) is the extracellular matrix (ECM) within which cumulus cells are embedded. Studies using animal models support the roles of adequate cumulus oocyte complex (COC) matrix in not only ovulation but also fertilization (Zhuo and Kimata, 2001; Russell and Salustri, 2006). Another point of clinical relevance to cumulus cells that merits mention is the routine removal of these cells for intra cytoplasmic sperm injection (ICSI) within a 2–4 h window post-retrieval. Given the known influences of cumulus cells on oocyte maturation and fertilization, it is not unreasonable that a premature removal of cumulus cells may compromise the last steps of oocyte development and competence acquisition. For instance, oocytes that progressed from metaphase- I to metaphase-II shortly after retrieval exhibit compromised fertilization rates when compared to oocytes already in metaphase-II (De Vos et al., 1999). Extending the time of culture prior to intra cytoplasmic sperm injection (ICSI) may permit some of the oocytes to complete their developmental programs; a situation particularly relevant to those remaining slightly immature at the time of retrieval. For this subset of oocytes, studies tested the effects of delayed sperm injection on oocyte maturation and fertilization rates. The use of such a rescue maturation step was shown to provide an increased number of available embryos, albeit of compromised quality (Vanhoutte et al., 2005).

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1.2.2 Oocyte maturation stage The removal of the cumulus–corona cell mass gives a unique opportunity to evaluate oocyte morphology prior to fertilization, in particular, the nuclear maturation stage. Oocyte nuclear maturity, as assessed by light microscopy, is assumed to be at the MII stage when the first polar body is visible in the perivitilline space. The MII stage is characterized by the alignment of the homologous chromosomes on the spindle equatorial plate during metaphase of the second meiotic division. It is generally recognized that 85% of the retrieved oocytes following ovarian hyper stimulation display the first polar body and are classified as MII, whereas 10% present an intra-cytoplasmic nucleus called the ‘germinal vesicle’ characteristic of prophase I of the first meiotic division. Approximately 5% of the oocytes have neither a visible GV nor first polar body and these oocytes are generally classified as MI oocytes (Rienzi and Ubaldi, 2009). Additional information on oocyte nuclear status can be obtained with the use of polarized light microscopy combined with image processing software for the non-invasive visualization of the meiotic spindle and other oocyte birefringent structures. The meiotic spindle is a microtubular structure involved in chromosome segregation; therefore, it is crucial in the sequence of events leading to the correct completion of meiosis and subsequent fertilization. Parallel-aligned meiotic spindle microtubules are birefringent and able to shift the plane of polarized light inducing retardance. These properties enable the system to generate contrast and image the meiotic spindle (MS) structure (Oldenbourg, 1999).

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The presence of the meiotic spindle (MS) gives more accurate information about the nuclear stage of the oocyte. In particular, some oocytes can be immature (at the stage of early telophase I) when observed with polarized light microscopy, despite the presence of first polar body in the perivitilline space. At this stage, in fact, there is continuity between the ooplasm of the oocyte and the forming first polar body (PBI) and the meiotic spindle (MS) is interposed between the two separating cells. This condition normally lasts for 75–90 min. The meiotic spindle MS has been found to disappear in late telophase I, reforming only 40– 60 min later (Montag et al., 2011). However, it must be underlined that other factors, such as sub-optimal culture conditions, temperature fluctuations and chemical stress during manipulation, can contribute to meiotic spindle( MS) disassembly (Rienzi and Ubaldi, 2009). Finally, the percentage of oocytes with detectable meiotic spindle (MS) is also related to the time elapsed from hCG administration and is higher after 38 h (Cohen et al., 2004).

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1.2.3 Oocyte size and shape A critical oocyte size is necessary for resumption of meiosis (Otoi et al., 2000). At the beginning of oocyte growth, size is determined by strong adhesion between the oolemma and the inner zona surface (Tartia et al., 2009).The mean ovarian diameter of MII oocytes may vary substantially but it is not related to fertilization or developmental quality of human ICSI embryos at the cleavage stage of development (Roma˜o et al., 2010). The situation is different with giant oocytes (Rosenbusch et al., 2002) this type of oocyte has about twice the volume of a normal oocyte (about 200 µm) and is tetraploid before meiosis due to their origin, i.e. nuclear but no cytoplasmic division in an oogonium or cytoplasmic fusion of two oogonia. These mechanisms explain the binucleate appearance of prophase I giant eggs. These oocytes always contribute to digynic triploidy and must never be transferred, although the presence of at least one giant oocyte in a cohort of retrieved eggs has no effect on treatment outcome (Machtinger et al., 2011). It is evident that oocytes with extreme forms of shape anomaly exist (Esfandiari et al., 2005) such ova have been shown to be fertilizable and may lead to the birth of healthy babies. While quantifying the degree of the elongation, some authors (Ebner et al., 2008) realized that the dimensions of the shape anomaly were neither correlated with fertilization nor embryo quality. However, when oocytes with ovoid zonae develop, day 2 embryos show a flat array of blastomeres rather than the more traditional tetrahedral arrangement and further development is often delayed (Ebner et al., 2008). Rarely, two oocytes can be found within one follicular complex. Each oocyte is usually surrounded by a zona pellucida but the ZP immediately between the two oocytes is commonly shared rather than duplicated. It is not 20

uncommon for these conjoined oocytes to show different nuclear maturational states. It has been suggested that such oocytes may play a role in producing dizygotic twins; however, even when both of the conjoined oocytes are mature it is rare that both fertilize and no pregnancies have been reported from such oocytes (Rosenbusch and Hancke, 2012). 1.2.4 Cytoplasmic features It has been shown in the literature that severe dysmorphisms of the cytoplasmic texture impairs the developmental and implantation potential of the embryo (Balaban and Urman, 2006). Although a homogeneous, normal cytoplasm is expected in recovered oocytes the biological significance of different degrees of heterogeneity in the ooplasm is unknown (Rienzi et al., 2012). Based on current evidence, slightly heterogeneous cytoplasm may only represent normal variability among retrieved oocytes rather than being an abnormality of developmental significance (Alpha Scientists in Reproductive Medicine and ESHRE Special Interest Group of Embryology, 2011). Granularities of the cytoplasm are poorly defined in the literature and may depend on the modulation of the optical path used in phase contrast microscopy. Granular cytoplasm and in particular centrally located granular area was associated with poorer pronuclear stage and embryo quality. These cytoplasmic alteration may be a sign of oocyte immaturity (Kahraman et al., 2000; Cota et al., 2012) These morphological deviations should be carefully distinguished from inclusions such as refractile bodies or lipofuscin bodies (Otsuki et al., 2007) and organelle clustering which is also described as very condensed centralized granularity detectable by any form of microscopy (Alpha Scientists in 21

Reproductive Medicine and ESHRE Special Interest Group of Embryology, 2011). One of the most important severe cytoplasmic abnormalities of MII oocytes is the appearance of smooth endoplasmic reticulum (SER) clusters (SER discs) within the cytoplasm. Smooth endoplasmic reticulum (SER) discs can be identified as translucent vacuole-like structures in the cytoplasm by phase contrast microscopy. Evidencebased data clearly demonstrates that embryos derived from oocytes with SER discs are associated with the risk of serious, significantly abnormal outcomes (Ebner et al., 2006). Vacuoles within the cytoplasm are defined as fluid-filled structures which can be more easily noticeable and differ from SER discs. It is believed that vacuoles arise either spontaneously or by fusion of preexisting vesicles derived from SER and / or Golgi apparatus (El Shafie et al., 2000; Reinzi et al., 2012). Small vacuoles of 5–10 µm in diameter are unlikely to have a biological consequence, whereas large vacuoles >14 µm are associated with fertilization failure. How vacuolization affects pronuclear formation is unclear; probably, the presence of these cytoplasmic inclusions interfere with cytoskeleton function (Ebner et al., 2006). A negative effect of these structure on MII meiotic spindle has also been suggested (Van Blerkom et al., 1990). In oocytes that are fertilized, those vacuoles that persist beyond syngamy can interfere with cleavage planes, resulting in a lower blastocyst rate (Alpha Scientists in Reproductive Medicine and ESHRE Special Interest Group of Embryology, 2011).

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1.2.5 Extra cytoplasmic features 1.2.5.1. Zona pellucida The zona pellucida (ZP) is the specialized electron microscopy layer that directly surrounds the oocyte. As such, the ZP represents the interface between the oocyte and its enclosing cumulus cells. The normal development of follicles, and thus the oocytes, depends on the presence and integrity of the ZP (Rankin et al., 2001). Involved in oocyte–somatic cell interactions, the ZP also plays essential roles at fertilization permitting sperm–egg interactions, the acrosome reaction, and an adequate block to polyspermy. The clinical introduction of intra cytoplasmic sperm injection (ICSI) has permitted not only the treatment of male factor infertility and the ability to circumvent many cases of fertilization failures but also the identification of variants in ZP morphology. In contrast to IVF, the removal of cumulus cells before ICSI allows the visualization of the ZP shortly after retrieval and prior to fertilization. A large number of ZP variants (appearance, thickness, irregularities, composition, and organization) have been described with the advent of ICSI. Thicker ZPs are associated with decreased fertilization rates, implantation, and pregnancy rates (Bertrand et al., 1995). The dynamic features of the ZP were also later evaluated; with thickness variation (ZPTV) measured along the circumference of the zona; the more variable the ZP thickness, the better the quality of the embryos (Host et al., 2002), and the higher the implantation and pregnancy rates (Gabrielsen et al., 2000). During oogenesis, the ZP is laid down in a very orderly fashion (in both time and space). Given its critical role during oocyte and embryo development, the zona is likely a good predictor of oocyte quality. The ZP has been imaged at different time 23

points, on the day of retrieval or post embryo culture. While ZP thickness varies and becomes thinner post fertilization, there is still a clear value of examining the zona on the day of retrieval, that is after it has reached its maximal thickness post oocyte maturation and before later remodeling events. Interestingly, in the clinic the organization of the ZP appears influenced by intrinsic factors in the oocyte from the time of retrieval, hormonal stimulation (Bertrand et al., 1996) and subsequent ZP thickening and hardening during culture. The ZP also plays a pivotal role in pre implantation embryos; for instance, abnormalities in oocyte (and thus ZP) shape are associated with irregular cleavage patterns, compromised

cell–cell

contacts,

and

subsequent

difficulties

in

developmental progression (Ebner et al., 2008). The importance of the ZP continues until the blastocyst stage, a time when the embryo needs to hatch out of the zona prior to implanting into the uterine epithelium. Interestingly, the zona pellucida thickness may reflect a subsequent ability of the embryo to hatch and implant successfully (Gabrielsen et al., 2000). Although still contentious, there may be a causal relationship between recurrent implantation failure and an inability of embryos to escape normally from the ZP (Edi-Osagie et al., 2003). Since apparent changes in thickness or complete absence of the ZP are extremely rare (Stanger et al., 2001); the degree to which discoloration of the ZP contributes to the birefringence of the outer shell is not known. However, it has been suggested that successful fertilization, embryo development and pregnancy can be achieved after transfer of embryos derived from brown zonae (Esfandiari et al., 2006).

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b. Perivitelline space The perivitelline space (PVS) represents the acellular compartment in between the plasma membrane of the oocyte and it’s ZP. It becomes clearly visible in a mature oocyte with the extruded polar body located in its most prominent portion. There is a gradient of possible PVS arrangements: from directly opposed to the membrane, to separate and distinguishable, to be exaggerated. These variants may exist within the circumference of a single female gamete or across a cohort of oocytes (Rawe and Combelles, 2009). Several authors have noted that approximately one-third of all ova show a large PVS, a dysmorphism that was found to be negatively correlated with fertilization rate and embryo quality (Xia, 1997; Rienzi et al., 2008). Data from the literature indicate that a large PVS may be ascribed to over-mature eggs (Miao et al., 2009); that is, such eggs have shrunk in relation to zona pellucida presenting a large gap between the oocyte and the surrounding zona. A large PVS also occurs if a larger portion of cytoplasm is extruded together with the haploid chromosomal set during first polar body formation. This would result in a large first polar body and as a consequence a large PVS (Rienzi et al., 2012). One retrospective study demonstrated a correlation between a large PVS and a decline in both fertilization and embryo quality (Xia, 1997), while others reported a lack of any such relationships (Plachot et al., 2002). Recently, Ten et al., (2007) reported no influence of PVS size on fertilization, but improved embryo quality in oocytes with large PVS. Relatively little attention has been given to the composition of the PVS, although it does contain a matrix of proteins and hyaluronic acid organized as granules and filaments (Dandekar and Talbot, 1992). The exact origin of the PVS chemicals remains unknown, although the 25

cumulus cells and oocyte represent likely candidates. Indeed, there is some support for the presence of cortical granule material in the PVS, as well as the formation of a cortical granule envelope within the PVS (Talbot and Dandekar, 2003). Another measurable morphological feature of the PVS is the presence or absence of debris, with about 40% of donor oocyte (Ten et al., 2007) and 50% of zygote cohorts (Plachot et al., 2002) displaying PVS debris. The origin of such debris, as observed in the clinic, remains unknown. Xia (1997) proposed that it might arise from premature exocytosis of cortical granules. c. Polar body The mammalian oocyte is arrested in diakinesis of prophase I at birth. It retains an intact nucleus (the germinal vesicle, GV) until ovulation at which time exposure to luteinizing hormone (LH) triggers meiotic resumption. After germinal vesicle breakdown (GVBD), a short anaphase I (A-I) and telophase I (T-I) take place and the first polar body (PB) is extruded. Finally, the oocyte becomes arrested at the metaphase-II (MII) stage until fertilization (Rawe and Combelles, 2009). Generally, first polar body morphology can be seen as a reflection of postovulatory age of the oocytes since this by-product of meiosis fragments with time. Nevertheless, the impact of first polar body morphology on outcome is still a matter of debate (Rienzi et al., 2012). Oocytes showing an intact PBI give rise to higher rates of implantation and pregnancy (Ebner et al., 1999) probably due to an increase in blastocyst formation (Ebner et al., 2002). Apparently, the benefit of selecting oocytes according to the morphology of PBI is diluted with increasing time between ovulation induction and ICSI, since studies with different schedules could not find a relationship between morphology of 26

PBI and subsequent ICSI outcome (De Santis et al., 2005, Fancsovits et al., 2006). However, the fact that a large PBI has a very poor prognosis remains unchallenged (Fancsovits et al., 2006). When large PBI’s are extruded, embryos with multinucleated blastomeres are a significantly more frequent consequence than for all other PBI morphological categories (Fancsovits et al., 2006). It has been postulated that the extrusion of an abnormally large PBI is due to dislocation of the MS (Verlhac et al., 2000). Sometimes it is difficult to distinguish between heavily fragmented PBI’s and debris within the perivitilline space. Fertilization and cleavage rate as well as embryo quality have been found to be unaffected by the presence of coarse granules in the PVS, however, the rates of implantation and pregnancy seem to be influenced (Farhi et al., 2002). Granularity in the PVS has been associated with over-maturity of oocytes (Miao et al., 2009).

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1.3 Ovulation Ovulation is the rupture of follicle at the surface of the ovary, releasing the oocyte. Ovulation has been likened to an inflammatory response, a description that remains highly germane (Espey, 1980; Espey and Lipner, 1994). To prepare for ovulation, the ovary must undergo a series of closely regulated events. Small follicles must mature to the preovulatory stage, during which the oocyte, granulosa cells, and theca cells acquire specific functional characteristics. The oocyte becomes competent to undergo meiosis, granulosa cells acquire the ability to produce estrogens and respond to luteinizing hormone (LH) via the LH receptor, and theca cells begin to synthesize increasing amounts of androgens that serve as substrates for the aromatase enzyme in the granulosa cells (Eppig, 1991; Richards, 1994). In response to the pre-ovulatory LH surge, the germinal vesicle of the oocytes in pre-ovulatory follicles undergoes germinal vesicle breakdown (GVBD), followed by first polar body extrusion (Kawamura et al., 2011). In the days immediately preceding ovulation, under the influence of FSH and LH, the secondary follicle grows rapidly to a diameter of 25 mm. Coincident with final development of the secondary follicle, there is an abrupt increase in LH that causes the primary oocyte to complete meiosis I and the follicle to enter the pre ovulatory stage. Meiosis II is also initiated, but the oocyte is arrested in metaphase approximately 3 hours before ovulation, in the meantime the surface of the ovary begins to bulge locally, and at the apex an avascular spot, the stigma, appears. The high concentration of LH increases collagenase activity, resulting in digestion of collagen fibers surrounding the follicle. Prostaglandin levels also increases in response to the LH 28

surge and cause local muscular contraction in the ovarian wall. Those contractions extrude the oocyte, which together with its surrounding granulosa cells from the region of the cumulus oophorus, breaks free (ovulation) and floats out of the ovary. Some of the cumulus oophorus cells then rearrange themselves around the zona pellucid to form the corona radiate (Sadler, 2006). Human chorionic gonadotrophin (hCG) has been the gold standard for ovulation induction as a surrogate for the midcycle LH surge for several decades. Due to structural and biological similarities, hCG and LH bind to and activate the same receptor, the LH/hCG receptor (Kessler et al., 1979). An important difference, however, exists between the half-life of LH and hCG, as the half-life of LH is approximately 60 min (Yen et al., 1968) whereas that of hCG is >24 h (Damewood et al., 1989). Due to its prolonged circulatory half-life, hCG exerts a sustained luteotropic activity, and may induce the occurrence of ovarian hyper stimulation syndrome (OHSS) (Delvigne and Rozenberg, 2002). Moreover, studies have reported adverse effects of hCG in terms of a reduced endometrial receptivity and a negative impact on oocyte quality (Forman et al., 1988; Valbuena et al., 2001). The role of the mid-cycle FSH surge in the natural cycle is not fully understood, but FSH has been shown to induce LH receptor formation in the luteinizing granulosa cells, thus optimizing the function of the corpus luteum. Moreover, FSH specifically seems to promote oocyte nuclear maturation, i.e. resumption of meiosis (Yding Andersen et al., 1999) and cumulus expansion (Stickland and Beers, 1976; Eppig, 1979). Interestingly, several studies reported the retrieval of more mature oocytes after GnRHa trigger, which could be an effect of a more physiological surge including a FSH surge as well as an LH surge (Humaidan et al., 2010; Oktay et al., 29

2010). Importantly, although large-scale studies do not exist, small studies have suggested that triggering of ovulation with GnRHa prevents ovarian hyper stimulation syndrome (Orvieto, 2005; Griesinger et al., 2007a; Hernandez et al., 2009)

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1.4 Fertilization A series of dynamic and complex events are triggered following sperm– oocyte interaction that sequentially leads to fertilization and the formation of zygote. These events include sperm penetration, sperm–oocyte fusion and oocyte activation, male and female pronucleus (PN) development and gradual migration of the pronuclei (PNs) to a central position in the oocyte (Papale et al., 2012). The fertilized oocyte undergoes maternal-tozygotic transition as a result of major changes in the molecular signals that control the arrest of meiotic development at the metaphase II stage of the second meiotic division (Ajduk et al., 2011). In humans, the sperm centriole has the leading role in organizing the microtubules, which direct the migration of PNs and their rotation within the cytoplasm. In this way, pronuclei (PNs) position their axis toward the second polar body and achieve a proper orientation at syngamy by controlling the plane of the first mitotic division (Papale et al., 2012). During pronuclei (PN) formation, nuclear precursor bodies (NPBs) become evident and start to migrate and merge into nucleoli in a time-dependent manner. The NPBs do not form a functionally active nucleolus at the zygote stage; however, they can be used as an indirect measure of the location and the grade of condensation of DNA within the PNs. Nucleoli are the sites of synthesis of pre-rRNA and its availability is extremely important since newly synthesized rRNA is necessary for the translational processes when the embryonic genome becomes active (Gianaroli et al., 2003).

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Once pronuclei (PNs) are aligned onto a polar axis, parental chromosomes then separate in preparation for mitosis. The human zygote’s mitotic potential is paternally inherited with the spermatozoon delivering the centrosome (Sathananthan et al., 1996). A zygote’s morphological characteristics are accepted to be an inherent indicator of both gamete quality and subsequent embryo implantation potential (Alpha Scientists in Reproductive medicine and ESHRE Special Interest Group of Embryology, 2011). Many studies have underlined the predictive value of zygote morphological assessment through correlations with chromosome constitution and the incidence of zygotic arrest (Edirisinghe et al., 2005; Zamora et al., 2011). Recent strategies in embryo selection include sequential morphology assessment where pronuclei (PN) scoring has been shown to play an important role as an indicator of gamete constitution as well as a prognostic tool for embryo competence. Scoring of PNs has also proved to be useful in countries where restrictive legislation mandates selection at the zygote stage for embryo transfer and consecutive elimination or cryopreservation of sibling zygotes (Senn et al., 2006). Although numerous studies have associated positive clinical results with the implementation of pronuclei (PN) scoring, other reports have questioned the predictive value of pronuclei (PN) scoring systems and see no benefits or improvement in the outcome (Nicoli et al., 2010; Weitzman et al., 2010).

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Fertilization check is to be performed at 17±1 h post-insemination which may establish uniformity in the future and eliminate the variability in pronuclei (PN) scoring regimens that have confounded comparative analyses. It must be noted; however, that the processes associated with fertilization by conventional insemination lags 1 h behind fertilization using intra cytoplasmic sperm injection ICSI (Nagy et al., 1998). A more complete elucidation of events during the zygote stage, however, can be expected with the application of continuous monitoring through the introduction of time-lapse imaging instead of the traditional isolated observations using light microscopy (Montag et al., 2011). Normal fertilization is assessed by the presence of two centrally positioned, juxtaposed PNs with clearly defined membranes and two polar bodies. If an abnormal pronuclei ( PN) number is observed whether it be 1, or 3 or more, pronucleous PNs, a low viable pregnancy is to be expected thus the transfer of these zygotes is to be avoided (Reichman et al., 2010). Aberrant pronuclei (PN) size and position have been correlated with developmental arrest and aneuploidy and are represented by PNs of unequal size (>4 µm), localized far apart or peripherally or with the presence of fragmented or additional micronuclei (Scott et al., 2007). Correct alignment of pronuclei (PNs) onto the polar axis is considered a fundamental feature for the completion of the first cleavage division and normal sequential development (Scott, 2001). The presence of unequal number and size of NPBs aligned at the PN junction has been correlated with increased embryo competence (Tesarik et al., 2000; Scott, 2003). Cell cycle-related dynamics of pronuclei (PN) events have led many authors to investigate and correlate the presence, pattern and number of polar body to embryo developmental potential. As shown for mitotic 33

cells, the inequality in number, size or distribution of N polar body is correlated with abnormal development (Pedersen, 1998). Panel experts from the consensus workshop established three categories for pronuclei (PN) scoring that is based on the morphology of NPBs and PNs, namely: (i) symmetrical, (ii) non-symmetrical and (iii) abnormal (Alpha Scientists in Reproductive Medicine and ESHRE Special Interest Groupof Embryology, 2011). Category 1 includes zygotes presenting with equal numbers and size of NPBs, either aligned at the junction between PNs or scattered in both PNs. Category 2, non-symmetrical, comprises all other patterns including peripherally localized PNs. Category 3, abnormal, includes single NPB (‘bull’s eye’) or total absence of NPBs. The latter are found to be correlated with imprinting errors and delayed onset of functional NPBs and nucleoli formation in animal models (Svarcova et al., 2009). Reports on early cleavage checks have been demonstrated to be a beneficial tool in selecting embryos with high implantation potential and decreased chromosomal anomalies (Lundin et al., 2001).

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1.5 The cleavage stage embryo A precise embryo quality evaluation is of paramount importance to sustain a successful in vitro fertilization (IVF) program. In most IVF clinics around the world, this quality assessment relies mainly on the morphological evaluation of cleavage stage embryos. Embryologists should be able to correlates the features observed at the optical microscope with the implantation potential of each particular embryo (Ebner et al., 2003; Alpha Scientists in Reproductive Medicine and ESHRE Special Interest Group of Embryology, 2011). To achieve this goal, many scoring systems based on the morphological features of the dividing embryo have been developed (Holte et al., 2007). These embryo classification systems are based on the evaluation of the number of blastomeres, the degree of fragmentation, the symmetry of the blastomeres, the presence of multinucleation and the compaction status. It is very important that the features related to implantation potential are assessed accurately and similarly (Prados et al., 2012). Cleavage stage embryos range from the 2-cell stage to the compacted morula composed of 8–16 cells. The number of blastomeres is used as the main characteristic with the highest predictive value (Fisch et al., 2001). Good quality embryos must exhibit appropriate kinetics and synchrony of division. In normal-developing embryos, cell division occurs every 18–20 h. Embryos dividing either too slow or too fast may have metabolic and/or chromosomal defects (Munne´, 2006; Magli et al., 2007).

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1.5.1 Cell numbers The developmental stage of an embryo, defined as the number of blastomeres on Days 1, 2 or 3 after insemination is an essential predictive factor for subsequent implantation and pregnancy rates (1 cell to > 10 cells). For assessment of embryo cleavage (numbers of blastomeres), the currently accepted observation schedule for optimal cleavage rates was defined at the Istanbul consensus workshop to be: Day 1 (26±1 h postICSI, 28±1 h post-IVF), 2-cells; Day 2 (44± 1 h), 4-cells and Day 3 (68±1 h), 8-cells (Alpha Scientists in Reproductive Medicine and ESHRE Special Interest Group of Embryology, 2011). Early cleavage i.e. the first mitosis occurring before 26±1 h (ICSI) and 28±1 h (IVF) respectively, has been shown to correlate with numbers of good quality embryos, blastocyst development and pregnancy rates (Fenwick et al., 2002). A number of studies have shown that the transfer of 4-cell embryos on Day 2 of culture results in significantly higher implantation and pregnancy rates compared with the transfer of embryos with either lower or higher cell numbers (Scott et al., 2007). Correspondingly, several studies have shown that for Day 3 transfers, implantation and live birth rates are positively correlated with an increase in cell number on Day 3, with the 8-cell stage (having been a 4-cell embryo on Day 2) having the highest rates ( Racowsky et al., 2011). The cleavage stage of the embryo at the time of transfer also seems to have a role in predicting early pregnancy loss Hourvitz et al., (2006) found that five or less blastomeres in the best embryo transferred on Day 3 were correlated with early pregnancy loss. A correlation between cell numbers at distinct observation time points and chromosomal errors has also been reported. It was shown by Munne´ (2006) that Day 2 embryos with 4 cells had the lowest rate of 36

chromosomal errors, while Magli et al., (2007) showed the same to be true for embryos with 7- to 8 cells on Day 3. The same pattern was observed by Finn et al.,(2010) who described a higher rate of euploidy in embryos with seven to eight blastomeres on Day 3 compared with both six or less than six blastomeres and nine or more than nine blastomeres. 1.5.2 Fragmentation Small portions of cytoplasm enclosed by a cell membrane, but usually not containing DNA, are often formed during cell division. Fragmentation is, therefore, defined as the presence of anucleate structures of blastomeric origin (Keltz et al., 2006) and evaluation of the degree of fragmentation is included in almost every embryo scoring system. The degree of fragmentation is most often expressed as the percentage of the total cytoplasmic volume. The relative degree of fragmentation is defined as mild (<10%), moderate (10–25%) and severe (> 25%). It is often difficult to make a distinction between a large anucleate fragment and a small (nucleated) cell (Prados et al., 2012). Johansson et al., (2003) showed that portions of cytoplasm that were < 45 µm in diameter on Day 2 and < 40 µm in diameter on Day 3 did not contain DNA. It has been shown that a high degree of fragmentation correlates negatively with implantation and pregnancy rates (Racowsky et al., 2000), while the presence of minor amounts of fragmentation has no negative or possibly even a positive impact (Alikani et al., 1999). Increasing fragmentation also results in reduced blastocyst formation and can influence allocation of cells during differentiation (Hardy et al., 2003). The spatial distribution of the fragments in the perivitelline space (PVS) can be differentiated into two patterns: scattered and concentrated. The scattered appearance is found to correlate with an increased incidence of chromosomal abnormality (Magli 37

et al., 2007). The higher the degree of fragmentation the more difficult it is to differentiate between scattered and concentrated fragmentation. Fragmentation is considered to be an essential parameter to include in the evaluation of developing embryos, as embryos with very strong and persistent fragmentation are less likely to be viable (Prados et al., 2012). 1.5.3 Blastomere size It has been shown that a high degree of regularity in the blastomere size in embryos on Day 2 correlates with increased pregnancy outcome following assisted reproduction treatments (Holte et al., 2007). Uneven cleavage, i.e. a cell cleaving into two unequal sized cells, may result in an uneven distribution of cytoplasmic molecules, e.g. proteins and mRNAs, and has been shown to correlate with a higher incidence of multi nucleation and aneuploidy ( Magli et al., 2001). The relative blastomere size in the embryo is dependent on both the cleavage stage and the regularity of each cleavage division. The blastomeres of 2-, 4- and 8-cell embryos should be equal rather than unequal in size. In contrast, blastomeres of embryos with cell numbers other than 2, 4 and 8 should have different sizes as there is an asynchrony in the division of one or more blastomeres (Prados et al., 2012).

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1.5.4 Nucleation The nucleation status is defined as the presence or absence of nuclei in the blastomeres of the cleavage stage embryo. Ideally, the nucleation status of each blastomere in the embryo should be evaluated as a single nucleus per blastomere no nuclei visible or multinucleation. The most studied nucleation status is multinucleation, which is defined as the presence of more than one nucleus in at least one blastomere of the embryo (Van Royen et al., 2003). Multinucleation can be evaluated both in the early cleaved Day 1 (26–28±1 h post-insemination), Day 2 (44±1 h post - insemination) and Day 3 (68±1 h post-insemination) cleavage stage embryos; although the assessment of a Day 3 embryo may be more complicated due to the smaller cell size and the larger number of cells (Van Royen et al., 2003). Embryo quality has been shown to correlate with multinucleation, and 4-cell embryos on Day 2 and 8-cell embryos on Day 3 show reduced multinucleation compared with the other cell stages observed on these days (Ziebe et al., 2003). Multinucleation is predictive of a decreased implantation potential (Moriwaki et al., 2004) and multinucleated embryos are associated with an increased level of chromosome abnormalities (Agerholm et al., 2008) as well as an increased risk of spontaneous abortion (Scott et al., 2007). Multinucleation is more frequent in blastomeres originating from embryos with uneven cleavage compared with embryos with evenly cleaved blastomeres (Hardarson et al., 2001). Multinucleation can also be divided into binucleation (two nuclei per cell) or multi/micro nucleation (more than two nuclei per cell). These appearances probably have different origins (Meriano et al., 2004). Multinucleated embryos are usually excluded from transfer. However, it has been shown that 39

binucleated cells on Day 1 can cleave into chromosomally normal cells (Staessen and Van Steirteghem, 1998). On the other hand, severe multi nucleation is probably not compatible with normal cell cleavage. Multi nucleation evaluation should be included in any embryo assessment protocol to select the highest quality embryo for transfer. Although these embryos do give rise to live births, they should be excluded from selection for embryo transfer if an alternative embryo is available. The absence or presence of a single nucleus per blastomere has been shown to be a predictor of embryo implantation potential (Saldeen and Sundstro¨m, 2005).Visualization of four mono nucleated blastomeres in a 4-cell embryo predicted a higher implantation rate than in cases where zero to three mono nucleated blastomeres was seen (Saldeen and Sundstro¨m, 2005). However, other studies have found that grading embryo nuclear score on Day 2 had no additive value for the prediction of implantation rate above that predicted by Day 3 embryo morphology (Bar-Yoseph et al., 2011).

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1.6 Implantation Embryo implantation represents a critical step of the reproductive process and consists of a unique biological phenomenon. The blastocyst comes into intimate contact with the endometrium and forms the placenta that will provide an interface between the growing fetus and the maternal circulation (Guzeloglu-Kayisli et al., 2009). Successful implantation requires a receptive endometrium, a functional embryo at the blastocyst developmental stage and a synchronized dialog between maternal and embryonic tissues (Simon et al., 2000). The endometrium is a multilayered, dynamic organ overlaying the myometrium and comprises a functional layer and a basal layer. Each month, cells in the functional layer are separated from the basal layer during menstruation. The basal layer is attached to the myometrium and remains intact during menstruation, serving as a base for endometrial regeneration. The endometrium is composed of several different cell types, including luminal and glandular epithelial cells, stroma with stromal fibroblastic cells, immune competent cells and blood vessels. The number, activity, structure and function of these cells change throughout the menstrual cycle and change again during pregnancy (Diedrich et al., 2007). The human endometrium undergoes a complex series of organized proliferative and secretory changes in each menstrual cycle, and exhibits only a short period of receptivity, known as the ‘window of implantation’ (Strowitzki et al., 2006). The endometrium becomes receptive to blastocyst implantation ~ 6 days after ovulation and remains receptive for 4 days (Bergh and Navot, 1992).

41

When implantation does not occur, a timely destruction of the fully developed endometrium leads to menstruation. However, if implantation occurs, the endometrium continues to grow and undergoes further morphological and molecular changes to provide sufficient support for the growing embryo (Strowitzki et al., 2006) 1.6.1 The proliferative phase The main feature of the endometrial tissue during the proliferative phase is active proliferation and angiogenesis to ensure nutrition of the developing new tissue, while suppressing apoptotic factors. Adequate development of endometrial tissue during this phase of the cycle is crucial for synchronization of the maturation process necessary for implantation during the secretory phase endometrium (Hoozemans et al., 2004). One of the key players during the proliferative phase is the rising female hormone estrogen (Strowitzki et al., 2006). In a highly proliferating tissue like the endometrium during the proliferative phase, angiogenesis is mandatory to supply the tissue with sufficient nutrients. Angiogenesis takes place throughout the menstrual cycle with a significant increase in the basal layer and in the sub epithelial plexus during the first part of the cycle. Angiogenesis changes in aspect during the cycle: major vessel elongation dominates the proliferative phase, whereas intussusception is the main mechanism during the early- to midsecretory phase (Gambino et al., 2002).

42

1.6.2 The secretory phase In humans, during the late secretory phase of normal menstrual cycles, the morphological changes that characterize decidualization are taking place independently of conception. All cell types in the endometrium are affected, with the functionally distinct tissue showing characteristic endometrial cell differentiation and an infiltration by large numbers of immune cells. Progesterone, which suppresses proliferation and induces cell differentiation, is the major player during this second half of the menstrual cycle, (Wang et al., 1998; Bergeron, 2000). 1.6.3 Endocrinological aspects Progesterone and estrogen are the dominant hormonal modulators of endometrial development. Ovarian estrogen and progesterone condition the uterus for implantation. Knowledge about the precise temporal action of these hormones within the menstrual cycle has allowed the development of hormone-based contraception. Both the epithelial and stromal compartments express progesterone and estrogen receptors and the response depend on the levels of these receptors as well as on the concentration of the hormones themselves (Diedrich et al., 2007).In addition to progesterone and estrogen, a number of other endocrinological factors are known to mediate endometrial function (Kodaman and Taylor, 2004). In rodents, prostaglandins (PGs) are thought to facilitate increased vascular permeability during implantation (Kennedy, 1979), as well as enzymes involved in PG production (COX-1 and COX-2) which show cyclical changes in expression (Chakraborty et al., 1996; Das et al., 1999).

43

Human chorionic gonadotrophin (hCG) is thought to have direct effects on the endometrium and also mediates cross-talk between the embryo and the endometrium, through chorionic gonadotrophin receptors present on epithelial cells (Srisuparp et al., 2003). 1.6.4 Apposition, adhesion and penetration Implantation has three stages: apposition, adhesion and penetration. Apposition is an unstable adhesion of the blastocyst to the endometrial surface. During this stage, the trophoblast becomes closely opposed to the luminal epithelium (Tabibzadeh and Babaknia, 1995). This is followed by the adhesion stage in which the association of the trophoblast and the luminal epithelium is sufficiently intimate as to resist dislocation of the blastocyst by flushing the uterine lumen. The first sign of the attachment reaction occurs on Day 20–21 in humans, and it coincides with a localized increase in the stromal vascular permeability at the site of blastocyst attachment (Sharkey and Smith, 2003). Following adhesion, the embryo invades through the luminal epithelium into the stroma to establish a relationship with the maternal vasculature. Although this activity is mainly controlled by trophoblasts, the decidua also limits the extent of invasion (Sharkey and Smith, 2003). In response to this invasion and the presence of progesterone stimulation, the endometrial stromal cells and endometrial extracellular matrix undergo decidualization that is essential for the viability of the pregnancy (Cakmak and Taylor, 2011).

44

Integrins are the best-studied group of cell adhesion molecules in the endometrium. They combine heterodimeric, non-covalently bound α and β subunits. Their extracellular domain serves as a receptor for various extracellular matrix (ECM) ligands such as fibronectin, collagen and laminin (Albelda and Buck, 1990). Osteopontin is an important receptor for the integrins and indeed a possible function for embryonic implantation seems obvious. Osteopontin has been detected during the mid- to late secretory phase (Apparao et al., 2001; von Wolff et al., 2001) in glandular epithelial cells and in uterine secretions from the secretory phase (von Wolff et al., 2001). LIF and LIF receptor in human endometrium were first described by Cullinan et al., (1996).The role of leukaemia-inhibitory factor (LIF) in implantation has been shown in a mouse model lacking a functional LIF gene; although the homozygous female mouse mated with a wild-type male the embryos did not attach and implant (Stewart, 1994). Data on humans, however, are still scarce. Clinically, more women with idiopathic infertility have undetectable levels of LIF in their uterine flushing in contrast with fertile women (Laird et al., 1997). The embryo expresses L-selectin after it hatches from the zona pellucida (Genbacev et al., 2003). L-selectin is expressed on vessel walls, where it captures leukocytes with their oligosaccaride structures and binds them after integrin activation at the site where they are needed (Alon and Feigelson, 2002).

45

Chapter two Patients and Methods

2. Patients and methods 2.1 Ethical consideration Informed consent was taken from all patients. All collected information from couples was kept confidential. This study was approved by the local scientific research committee on medical research and the ethical committee of the faculty of medical science in Sulaimani, University of Sulaimani, Kurdistan, Iraq. 2.2 Study design This study was carried out at Dwarozh- IVF center in sulaimani. A prospective analysis was performed on the data obtained from 132 patients undergoing ICSI cycle that had been performed between May 2013 and January 2014. All patients were referred to ICSI cycles for the following reasons: (i) female’s infertility factors (ii) unexplained infertility for more than 5 years. Male infertility factors were excluded. In this prospective study, morphological feature of 1056 metaphase II oocytes from 132 ICSI cycles were evaluated and compared. The rates of fertilization (two pronuclei were seen in the injected oocyte after 16-20 hours of ICSI), and high quality embryos (even, homogenous blastomeres with less than 15% fragmentation) were evaluated; besides implantation rates (total number of gestational sacs divided by total number of embryo transferred) were compared.

46

2.3 Ovarian stimulation The entire females’ partner had been screened at the 2nd day of their cycles for hormonal assay including FSH (3.5-12.5 mIU/ml), LH (2.412.6 mIU/ml), E2 (18-147 pg/ml), P4 (0.1-0.44 ng/ml) the screening was performed with (mini VIDAS, bioMerieux, 06267 M, France).

In

addition trans -vaginal ultrasonography (Siemens, Sonoline G20, Siemens Medical Solutions USA, Inc.) was performed for them in order to asses’ endometrial thickness, antral follicles count > 6 follicles and any underlining ovarian and uterine pathologies; all of which had been done by an expert gynecologist. In all patients short agonist protocols was used to stimulate follicular development. The short protocol includes pituitary desensitization with gonadotrphin releasing hormone agonist (Decapeptyl 0.1 mg, Ferring GmbH, Germany) at day 2 of the cycle and ovarian stimulation with follicle- stimulating hormone (fostimon, IBSA, Lugano 3, Suisse; Gonal- f, Laboroteries Serono S.A, Switzerland; Puregon, Schering-Plough, NV.Organon, Oss, Netherlands;) or combined LH and FSH (Merional, IBSA, Lugano3, Suisse; Menegon, Ferring GmbH, Germany) starting at day 3 of the cycle. Follicular development was monitored by measuring serum E2 and performing trans-vaginal ultrasound (Siemens, Sonoline G20, Siemens Medical Solutions USA). Ovulation and final maturation of the ova were induced with human chorionic gonadotrophin hCG (Choriomon, IBSA, Lugano 3, Suisse; Ovitrelle, Industria Farmacutica Serono S.P.A, Bari, Italy) as a single dose of 10 000 IU, when the leading follicle reached 18 mm in average diameter in addition to the presence of at least two other follicles of more than 16 mm in size and E2 > 500pg/ml then the oocytes were retrieved.

47

2.4 Oocyte retrieval Oocytes were aspirated 34- 36 hours after hCG administration. Oocyte retrieval was performed by Trans vaginal ultrasound guided puncture using 16 gage 35cm double lumen aspiration needle (William A.Cook, Australia pty Ltd.) as shown in plate (2.1) with negative pressure 20 mmHg under general anesthesia. Follicular fluid aspirated into tube containing aspiration media and heparin (Ferticult Aspiration, Fertipro N.V, Belgium), then the oocyte-cumulus complex was collected under stereomicroscope (Nikon, SMZ 1500, Tokyo, Japan) as shown in plate (2.2) and washed with IVF media (Ferticult IVF, Fertipro N.V, Belgium) then placed in a 4 well dish that contain IVF media (Ferticult IVF, Fertipro N.V, Belgium) in incubator 37˚C and 6 % CO2 for 1-2 hours. 2.5 Oocyte denudation Removal of the surrounding cumulus cells was accomplished by a combined enzymatic and mechanical treatment carried out under a stereoscopic dissecting microscope (Nikon, SMZ 1500, Tokyo, Japan). Denuding dish was prepared with a droplet of 100 µl of 80 IU/ml hyaluronidase enzyme in HEPES- buffered medium (Hyase, fertipro N.V, Belgium) and five droplets of IVF media ( Ferticult IVF, Fertipro N.V, Belgium) covered with mineral oil (Ferticult Mineral Oil fertipro N.V, Belgium), placed on the heated area in the hood to warm up for 10 minutes. Cumulus–oocyte complexes are transferred into the droplet of hyaluronidase solution and repeatedly aspirated through a Pasteur pipette for up to 30–40 seconds. At this time dissociation of the cells is initially observed. Further mechanical denudation is carried out in the enzymefree IVF media

(ferticult-IVF Fertipro N.V, Belgium) droplets by

repeated aspiration through commercially prepared stripper tips (Ez-strip, 48

Research Instruments Ltd, United Kingdom) with decreasing inner diameters of 290 and 135 µm. The oocytes are then transferred through the droplets of IVF media (ferticult-IVF, Fertipro N.V, Belgium) until all coronal cells have been finally removed and all traces of enzyme have been washed off. The denuded oocytes are placed in a culture dish that consist of nine small droplets of IVF media (ferticult-IVF, Fertipro N.V, Belgium) 5µl each that are arranged in a square of 3 x 3 within the dish covered by mineral oil (Ferticult Mineral Oil fertipro N.V, Belgium) and placed in incubator at 37˚C with 6%CO2 until the time of ICSI. 2.6 Oocyte evaluation Oocytes are assessed for their maturation and for their morphology under an inverted microscope (intgera Ti, R.I., Olympus, IX51/IX70, Tokyo, Japan) at 400X magnification as shown in plate (2.3). The MII stage is characterized by the alignment of the homologous chromosomes on the spindle equatorial plate during metaphase of the second meiotic division. It is generally recognized that retrieved oocytes following ovarian hyper stimulation and displaying the first polar body are classified as MII, whereas the presence of an intra-cytoplasmic nucleus called the ‘germinal vesicle’ characteristic of prophase I of the first meiotic division as shown in micrograph (3.1) while oocytes having neither a visible GV nor first polar body are generally classified as MI oocytes, (Reinzi and Ubaldi, 2009) as shown in micrograph (3.2).

49

The MII oocytes used for micromanipulation were grouped as follows: (1) Normal MII oocytes showed a clear cytoplasm with homogeneous fine granularity, a round or ovoid first polar body with smooth surface and size within a small perivitelline space, and a colorless zona pellucida with regular shape as shown in micrograph (3.3) (Ten et al., 2007). (2) Abnormal MII oocytes with cytoplasmic abnormalities such as (i) inclusion body as shown in micrograph (3.4). (ii) Presence of vacuoles, which

are

considered

fluid-filled

membrane-bound

cytoplasmic

inclusions, as shown in micrograph (3.5) and can be clearly distinguished morphologically from smooth endoplasmic reticulum clusters (sERC) under the inverted microscope. These clusters are not separated from the rest of the cytoplasm by a membrane and are implicated in implantation failure. Therefore, in this study; the presence of sERC was not evaluated. (iii) Granular and dark cytoplasm as shown in micrographs (3.6), (3.7), (3.9) and (3.13). (3) abnormal MII oocytes with extra cytoplasmic abnormalities such as large polar body as shown in micrographs ( 3.8 and 3.12), fragmented polar body as shown in micrograph (3.9), dark zona pellucida as shown in micrograph (3.10), wide perivitelline space as shown in micrographs (3.10 and 3.12) and perivitelline debris as shown in micrographs (3.11 and 3.13). (4) MII oocytes with combined cytoplasmic and extra cytoplasmic abnormalities as shown in micrograph (3.13). Shape abnormalities were not recorded separately.

50

2.7 Semen preparation The semen was collected by masturbation in all cycles; then allowed to be liquefied for 15-30 minutes. After liquefaction analysis was done with Makler chamber (Polymedco Inc., Yorktown, NY) using phase contrast microscope (Olumpus, BX42, Tokyo, Japan) according to WHO criteria. The semen was prepared using colloidal silica density gradient centrifuging (Sil-select Stock, Fertipro N.V, Belgium). A dual gradient system was prepared (45%-90%) by mixing 1ml Sil-select with 9ml flushing media(ferticult flushing media, Fertipro N.V, Belgium) for high density gradient 90%; and mixing of 5.5 ml of flushing media with 4.5 ml of Sil-select stock for low density gradient 45%. Then 2.5 ml of the high density gradient 90% was transferred into a sterile disposable centrifuge tube with sterile pipette; then 2.5 ml of density gradient 45% was slowly transferred over the high gradient layer then 2.5ml of liquefied semen was gently added onto the upper layer using a sterile pipette. After the preparations the layers were centrifuged with (Rotofix 32 A, Hettich, Germany) for 15-18 minutes at 2500 rpm. When centrifugation finished, the supernatant was removed down the pellet. The pellet was then mixed with 2-3 ml of flushing media and centrifuged for 8-10 minutes. Supernatant was removed and pellet was mixed again with 0.5ml of flushing media and kept in an incubator 37˚C until the time of ICSI.

51

2.8 Intra cytoplasmic sperm injection procedure An ICSI dish (Nunc A/S, Denmark) was performed by arranging three small droplets of IVF media (ferticult-IVF, Fertipro N.V, Belgium), 5µl each in this dish; these droplets are used for the denuded oocytes. A drop of the prepared sperm was placed in the ICSI dish. An additional droplet was placed in the middle of the dish containing 4µl of 10% polyvinyl pyrrolidone (10% PVP in Ferticult flushing medium, Fertipro N.V.,Belgium) which is a viscous medium for reducing sperm motility. The droplets were then covered with 5ml of mineral oil (Ferticult Mineral Oil fertipro N.V, Belgium) in order to maintain stable temperature, osmolality and PH (Miller et al., 1994). Active spermatozoa was aspirated by injection pipette (Injection pipette, Research Instruments Ltd, United Kingdom) at x 400 magnification and transferred to the droplet of the 10% PVP for immobilization. This was accomplished by gently lowering the tip of the pipette so as to compress the mid region of the sperm flagellum against the bottom of the dish, and then it was drawn into the injection pipette from its tail (Brooks et al., 2000). Oocytes were held on the special holding pipette (Holding pipette, Research Instruments Ltd, United Kingdom) with the polar body at the 6 or 12 o’clock position and the injection pipette (Injection pipette, Research Instruments Ltd, United Kingdom) which contained immobilized sperm was inserted into the ooplasm. ICSI was carried out on heated stage 37˚C of an inverted microscope (intgera Ti, R.I., Olympus, IX51/IX70, Tokyo, Japan) then the injected oocytes were washed and stored in a culture dish consisting of culture medium under mineral oil. They were kept in an incubator at 37˚C and 6% of CO2 as shown in plate (2.4).

52

2.9 Assessment of fertilization, embryo cleavage and embryo transfer Fertilization assessments were performed 17± 1 hour post injection. Normally, fertilized oocytes should be spherical and have two polar bodies and two PNs. PNs should be juxtaposed, approximately the same size, centrally positioned in the cytoplasm with two distinctly clear, visible membranes as shown in micrograph (3.14) (Papale et al., 2012). Embryo quality was evaluated under an inverted microscope (intgera Ti, R.I., Olympus, IX51/IX70, Tokyo, Japan). The following parameters were recorded: (1) the number of blastomeres; (2) the fragmentation percentage; (3) variation in blastomere symmetry (4) defects in the zona pellucida and the cytoplasm. High-quality (grade A) embryos were defined as those having all of the following characteristics: either 4–6 cells on day 2 or 8–10 cells on day 3 of development, less than 15% fragmentation, symmetric blastomeres, colourless cytoplasm with moderate granulation with no inclusions, absence of perivitelline space granularity and absence of zona pellucida dysmorphism, as shown in micrograph (3.15 and 3.17). Embryos lacking any of the above characteristics were considered as low quality (de Almeida et al., 2010), as shown in micrograph (3.16 and 3.18). For each couple, 1-4 embryos were transferred, depending on the embryo quality and the female’s age. Embryo transfer was cancelled if no embryos were available. Embryo transfer was performed on day 2 or day 3 using a Gynetics catheter (Gynetics medical products N.V, Lommel, Belgium). Transfers were performed with trans-abdominal ultrasound guidance. All women received 75 mg of aspirin tablet (Bristol) daily. The luteal phase was supplemented 53

with intra

vaginal micronized

progesterone (Uterogestan 100mg, Laboratoires Besins International-5 rue du Bourg I Abbe- Paris- France) at a dose of 600 mg daily in three divided doses starting on the day of oocyte retrieval and 250 mg of progesterone intramuscular (Primolute depot 250mg, Bayer Schering Pharma AG, Berlin, Germany) as a single dose every three day starting from the day of embryo transfer. Pregnancy was defined as one hCG titer assessed 16 day after embryo transfer. When positive, the first ultrasound scan was scheduled at 4 weeks after embryo transfer. Micrograph was taken with video camera 3mega pixel (WAT-221S, Olympus, Tokyo, Japan) loaded over the inverted microscope connected to Cronus software (Research Instruments Ltd, United kingdom). 2.10 statistical analysis Results are expressed as mean ± standard deviation (SD) for numeric variables and percentage for categorical variables. Mean values are compared by students t-test, and proportions are compared by z- test. The results are considered to be significant at the (P <0.05). Data analysis is carried out using the statistical analysis program (Statistical package for social science SPSS; version 20).

54

Plate (2.1) Double Lumen Aspiration Needle (William A. Cook, Australia pty Ltd.)

Plate (2.2) Nikon Stereo Microscope ( Nikon, SMz1500, Tokyo, Japan)

55

Plate (2.3) Inverted ICSI Microscope (integra Ti, R.I. Olympus, IX51/IX70, Tokyo, Japan)

Plate (2.4) CO2 incubator (Galaxy R Plus 170, NSB, UK)

56

Chapter three Results

3. Results 3.1 General characteristics: From the 132 patients included in this study, 56.81% were due to females- factors infertility while 43.18% were due to unexplained infertility. Mean age of patients was (33.27 ± 5.81) years. A total of 1200 oocytes were retrieved. Of these 1056 (88%) were mature in metaphase II stage and the remainder were immature oocytes either at metaphase I stage 84 (7%) or germinal vesicle stage 60 (5%). On assessment of metaphase II oocytes, 180 oocytes (17.04%) had normal MII oocytes morphology, 876 (82.95%) had at least one demonstrable morphological abnormality. Total injected MII oocytes were 954; of these 774 had abnormal oocytes morphology and 180 normal oocytes morphology. The number of embryo transferred was 363 with mean (2.77±1.40) as shown in Table (3.1).

57

Table (3.1): General Characteristics of ICSI cycles Characteristics

Data

Cycles (n)

132

Mean age ± SD (years)

33.27 ± 5.81

Oocyte retrieved (n)

1200

Number of MII oocytes (n)

1056

MII oocytes/total no. of oocyte retrieved (%)

(88)

Number of MI oocytes (n)

84

MI oocyte/total no. of oocyte retrieved

(7)

Number of oocytes at germinal vesicle stage (n)

60

Oocytes at GV stage/total no. of oocytes retrieved (%)

(5)

Number of normal MII oocytes (n)

180

Normal MII oocytes/total no. of MII oocytes (%)

(17.04)

Number of abnormal MII oocytes (n)

876

Abnormal MII oocytes/total no. of MII oocytes (%)

(82.95)

Total number of injected MII oocytes (n)

954

Total number of non-injected oocytes (n)

102

Total number of embryo transferred(mean ±SD)

363(2.77±1.40)

Values in parentheses are percentages , n = number of oocytes, MII = metaphase II, MI = metaphase I, GV = Germinal vesicle

58

From 876 abnormal MII oocytes morphology, Cytoplasmic abnormalities (dark cytoplasm, granular cytoplasm, presence of inclusion body and vacuoles) were seen in 516 (58.9%) of the abnormal MII oocytes. Extra cytoplasmic abnormalities (large /fragmented polar body, large and debris of perivitelline spaces) were seen in 104 (11.87%) while combined cytoplasmic and extra cytoplasmic abnormalities were responsible for the remaining 256 (29.22%) as shown in Table (3.2).

Table (3.2): Distribution of abnormal MII oocytes according to their morphology Characteristics

Data

No. of MII oocytes with cytoplasmic abnormalities(n)

516

MII oocytes with cytoplasmic abnormalities

(58.9)

/total no. of abnormal MII oocytes (%) No. of MII oocytes with extra cytoplasmic abnormalities (n)

104

MII oocytes with extra cytoplasmic abnormalities

(11.87)

/total no. of abnormal MII oocytes (%) No. of MII oocytes with combined cytoplasmic and

256

Extra cytoplasmic abnormalities (n) MII oocytes with combined abnormalities

(29.22)

/total no. of abnormal MII oocytes (%) Values in parentheses are percentages, n = number of oocytes, MII=metaphase II

59

3.2 Influence of MII Oocytes Morphology on Fertilization, cleavage and embryo quality rates From 876 abnormal MII oocyte morphology 774 oocyte were injected the rest of oocytes were not injected due to sever abnormalities while all 180 normal oocytes morphology were injected. The results show no significant differences in the fertilization, cleavage, and high quality embryo rates between the group of normal MII oocytes and the group of abnormal MII oocytes with (P value >0.05). Fertilization rates for morphologically normal and abnormal oocytes were (66.66% versus 60.2% respectively,). Although statistically not significant but the abnormal MII oocytes showed lower fertilization rate with P value >0.05 as shown in Table (3.3).

Table (3.3): Fertilization, Cleavage and Embryo quality rates after injection of normal and morphologically abnormal MII oocytes Normal MII

Abnormal MII P value

oocytes

oocyte

N=180

N=774

Fertilized oocytes (% per injected oocytes)

120 (66.66) 466 (60.2)

0.245

Cleaved embryos (% per fertilized oocytes)

113 (94.16) 440 (94.42)

0.914

High quality embryo (% per cleaved embryo)

66 (58.40)

0.812

264 (60)

Values in parentheses are percentages, N = number of injected MII oocytes

60

3.3 Comparisons between normal MII oocytes and MII oocytes with cytoplasmic abnormalities in relation to fertilization, cleavage and embryo quality From 516 oocytes with cytoplasmic abnormalities 454 oocytes were injected. The results show no significant differences in the fertilization, cleavage and high quality embryo of normal MII oocytes and MII oocytes with cytoplasmic abnormalities with P value >0.05. Fertilization rate between normal MII oocytes and MII oocytes with cytoplasmic abnormalities were (66.66% versus 58.81% respectively). Although statistically not significant but the oocytes with cytoplasmic abnormalities had the lowest fertilization rate as shown in Table (3.4).

Table (3.4): Compared results between normal MII oocytes morphology and MII oocytes with cytoplasmic abnormalities after injection Normal MII MII with cytoplasmic Oocytes

Abnormalities

N=180

N=454

Fertilized oocytes (% per injected 120 (66.66) 267 (58.81) oocytes) Cleaved embryos(% per fertilized 113 (94.16) 255 (95.50) oocytes) High quality embryos(% per cleaved 66 (58.40) 154 (60.39) embryos) Values in parentheses are percentages, N= number of injected MII oocytes

61

P value

0.1363 0.5675 0.7926

3.4 Comparisons between Normal MII oocytes and MII oocytes with extra cytoplasmic abnormalities in relation to fertilization, cleavage and embryo quality

From 104 oocytes with extra cytoplasmic abnormalities 89 oocytes were injected. Data from this study show that there was no significant difference in the fertilization, cleavage and high quality embryo rates between normal morphological MII oocytes and MII oocytes with extra cytoplasmic abnormalities with P value >0.05; however, high quality embryos were higher in the group with normal MII oocytes morphology than in the group with MII oocyte having extra cytoplasmic abnormalities as shown in Table (3.5).

Table (3.5): Compared results between normal MII oocytes morphology and MII oocytes with extra cytoplasmic abnormalities after injection Normal MII MII with extra

Fertilized oocytes)

oocytes

(% per

Oocytes

cytoplasmic abnormalities

N=180

N=89

P value

injected 120 (66.66)

59 (66.29)

0.9987

Cleaved embryos (% per fertilized 113 (94.16) oocytes)

55 (93.22)

0.8206

High quality embryos (% per cleaved 66 (58.40) embryos)

26 (47.27)

0.3333

Values in parentheses are percentages, N= number of injected MII oocytes

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3.5 Comparison between Normal MII oocytes and MII oocytes with combined abnormalities in relation to fertilization, cleavage and embryo quality From 256 oocytes with combined abnormalities 231 oocytes were injected. Data from this study show that there was no significant difference in the fertilization, cleavage and high quality embryo rates between normal morphological MII oocytes and MII oocytes with combined abnormalities with P value >0.05 as shown in Table (3.6).

Table (3.6): Compared results between normal MII oocytes and MII oocytes with combined abnormalities after injection Normal MII MII with combined

Fertilized oocytes)

oocytes

(% per

Oocytes

abnormalities

N=180

N=231

P value

injected 120 (66.66)

140 (60.60)

0.2861

Cleaved embryos (% per fertilized 113 (94.16) oocytes)

130 (92.85)

0.6807

84 (64.61)

0.4382

High quality embryos (% per cleaved embryos)

66 (58.40)

Values in parentheses are percentages, N= number of injected MII oocytes

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3.6 Relationship between MII oocytes morphology and implantation rate The result of this study show that there is a highly significant difference in implantation rate between the groups of normal MII oocytes morphology and abnormal MII oocytes morphology with p value <0.01 which was higher in the group of normal MII oocytes morphology (11.11%) than the MII oocytes with abnormal oocytes morphology (7.33%) as shown in Figure (3.1).

Figure (3.1): Copmarison of implantation rate between normal MII oocytes morphology and abnormal MII oocyte morphology

64

Data from this study show that there was highly significant difference in the implantation rate between normal MII oocytes and MII oocytes with cytoplasmic abnormalities with P value <0.01 which was higher in the group of normal MII oocytes (11.11%) than the group of MII oocytes with cytoplasmic abnormalities (9.03%) as shown in Figure (3.2).

Figure (3.2): Comparison of implantation rate between normal MII oocytes morphology and MII oocytes with cytoplasmic abnormalities

65

There was a highly significant difference in implantation rate between normal MII oocytes and MII oocytes with extra cytoplasmic abnormalities with P value <0.01 which was higher in the group of normal oocytes morphology (11.11%) than the MII oocytes with extra cytoplasmic abnormalities (2.38%) as shown in Figure (3.3).

Figure (3.3): Comparison of implantation rate between normal MII oocytes and MII oocytes with extra cytoplasmic abnormalities

66

There was a highly significant difference in implantation rate between normal MII oocytes and MII oocytes with combined abnormalities with P value <0.01 which was higher in the group of normal oocytes morphology (11.11%) than the MII oocytes with combined abnormalities (4.34%) as shown in Figure (3.4)

12%

11.11%

10% 8% 6%

4.34%

4% 2% 0% Normal oocyst Combined abnormality

Figure (3.4): Comparison of implantation rate between normal MII oocytes morphology and MII oocytes with combined abnormalities

67

GV

Micrograph (3.1) Immature oocyte at germinal vesicle (GV= germinal vesicle 400x)

Micrograph (3.2) Immature oocyte at Metaphase I stage (400x)

ZP

PB

PVS

Micrograph (3.1) Normal mature oocyte ay Metaphase II stage (PB=polar body, ZP=zona pellucida, PVS=perivittiline space 400x)

68

Micrograph (3.2) Abnormal MII oocytes with cytoplasmic inclusion body (200x)

Micrograph (3.3) Abnormal MII oocyte with cytoplasmic vacuoles (400x)

GV

Micrograph (3.4) Abnormal MII oocyte with central granulation of cytoplasm (400x)

69

Micrograph (3.5) Abnormal MII oocyte with granular cytoplasm (400x)

Micrograph (3.6) Abnormal MII oocyte with large and small polar body (200x)

Micrograph (3.7) Abnormal MII oocyte with fragmented polar body and central granulation (400x)

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Micrograph (3.8) Abnormal MII oocyte with dark zona pellucida large PVS (400x)

Micrograph (3.11) Abnormal MII oocyte with PVS debris (400x)

Micrograph (3.12) Abnormal MII oocyte with large polar body and large PVS (400x)

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Micrograph (3.13) Abnormal MII oocyte with dark cytoplasm and PVS debris (400x)

Micrograph (3.14) Zygote obtained by ICSI of oocyte displaying the 2 pronuclei (400x)

72

Micrograph (3.9) Day 2 high quality embryo (400x)

Micrograph (3.16) Day 2 low quality embryo (200x)

Micrograph (3.10) Day 3 high quality embryo (400x)

Micrograph (3.18) Day 3 low quality embryo (200x)

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Chapter four Discussion

4.1 Discussion As an attempt to improve the results of ICSI cycles, it is important to identify and utilize non- invasive parameters able to predict oocyte quality. Considering the vital role played by the oocyte in the developmental process, selection criteria involving the stage preceding fertilization would be extremely useful in selecting embryo for transfer (Setti et al., 2011; Cota et al., 2012). Nonetheless, previous reports are conflicting regarding the effects of oocyte morphological abnormalities on fertilization rate and embryo quality rate (Yakin et al., 2007; Patrizio et al., 2007; Setti et al., 2011). In this work, the presence of extra cytoplasmic and cytoplasmic oocytes dysmorphisms and their influence on fertilization, embryo quality and implantation rates in a total of 1056 MII oocytes, was evaluated. So far as is known, this is the first study in this regard carried out in Kurdistan Region of Iraq. The majority (60%-70%) of oocytes retrieved from stimulated cycles exhibited

one

or

more

abnormal

morphological

characteristics

(Mikkelsen and Lindenberg, 2001; Balaban and Urman, 2006; Ebner et al., 2006; Rienzi et al., 2008; Rienzi et al., 2012). The present study shows that 876 (82.95%) out of 1056 MII oocytes had at least one abnormal morphological characteristics. It is also observed that abnormal oocytes morphology was much higher than normal oocytes morphology 180 (17.04%). Similar results, though lower, was observed by (Balaban et al., 1998; Rienzi et al., 2008; Figueira et al., 2010) whose studies showed that > 60% of oocytes retrieved had at least one morphological abnormality, while a study by (Ten et al., 2007) found that 74

94.5% of all oocytes had at least one demonstrable anomaly. The best explanation for that was morphological variations of the oocyte may result from intrinsic factors such as age (de Bruin et al., 2004) and genetic defects, or extrinsic factors such as, the controlled ovarian stimulation protocol and the ovarian response to controlled ovarian stimulation (Rashidi et al., 2005, Ten et al., 2007; Figueira et al., 2010). The Result of current study clearly indicate that normal MII oocyte and abnormal MII oocytes morphology does not unfavorably alter the fertilization, cleavage and high quality embryo rates. Similarly, other studies (De Sutter et al., 1996; Balaban et al., 1998; Yakin et al., 2007; La Sala et al., 2009) demonstrated that the fertilization rate and embryo quality did not differ in the group of oocytes with no abnormality or one, two, or three morphological abnormalities. On the other hand studies by (Rienzi et al., 2008; Rienzi et al., 2011; Figueira et al., 2010) showed that oocytes dysmorphism is associated with a decreased fertilization, cleavage and embryo quality. Data from this study showed that there was a significant difference in implantation rate which was higher in the group with normal oocyte morphology 11.11% than the group with abnormal oocyte morphology 7.33% so our results agreed with results in Studies by (Meriano et al.2001; Rienzi et al., 2008) which established a significant relationship between MII oocytes morphology and implantation potential. Whereas Studies by (Balaban et al., 1998; Balaban and Urman, 2006; Yakin et al., 2007; La Sala et al., 2009) demonstrated that outcomes (implantation rate, pregnancy, take home baby and multiple pregnancy rates) were

75

similar when all embryos were derived from intact oocytes, or all from morphologically “handicapped” oocytes. In this study, Cytoplasmic and extra cytoplasmic abnormalities were observed in 58.9% and 11.87% respectively, indicating that among the abnormal oocytes morphology group, cytoplasmic abnormalities was much higher than extra cytoplasmic abnormalities. Similarly, a study by (De Sutter et al., 1996; Yakin et al., 2007; Rienzi et al,. 2008) revealed that extra cytoplasmic abnormalities were relatively lower than cytoplasmic abnormalities. In a study by Rienzi et al., (2008) extra cytoplasmic abnormalities were seen in 8.7% and cytoplasmic abnormalities were observed in 62%. The explanation for these cytoplasmic morphological abnormalities are probably multifactorial. Excessive ovarian response, characterized by an increased number of aspirated follicles and retrieved oocytes, has a detrimental effect on oocyte quality, resulting in a higher incidence of intra cytoplasmic defects. Deficient cytoplasmic maturity has been postulated to be reflected by cytoplasmic abnormalities (Kahraman et al., 2000). The oocytes retrieved from stimulated cycles may be derived from slowerdeveloping follicles, and therefore, the cytoplasm of these oocytes would be at different maturation stages upon the resumption of meiosis (Figueira et al., 2010; Setti et al., 2011). The result generated from our ICSI cycles demonstrated that cytoplasmic abnormalities had no effect on fertilization, cleavage, and embryo quality rates. Similarly results obtained from studies carried out by (De Sutter et al., 1996; Balaban et al., 1998; Balaban and Urman, 2006; Ten et al., 2007) show that cytoplasmic abnormalities did not affect the fertilization rate, embryo quality and clinical outcomes while The presence of 76

cytoplasmic features was correlated with impaired fertilization and embryo quality in studies by (Chamayou et al., 2006, Wilding et al., 2007; Rienzi et al., 2008; Figueira et al., 2010). In this study, implantation rate was statistically significant which was higher in the group with normal oocyte morphology 11.11% than the group with cytoplasmic abnormalities 9.03% similarly, results were obtained in studies by (Serhal et al., 1997; Wilding et al., 2007; Rienzi et al., 2008) showed that the presence of cytoplasmic features was correlated with lower implantation rate while studies by (Chamayou et al. 2006; Ten et al., 2007) have used a cumulative evaluation for cytoplasmic features of oocytes and the presence of these features did not influence pregnancy and implantation rates. In the present study, fertilization, cleavage and embryo quality was statistically not significant between the group of normal oocytes morphology and the group of oocytes with extra cytoplasmic abnormalities similarly studies by (Ebner et al., 2001; Plachot et al., 2002; Urman and Balaban, 2006; Yakin et al., 2007; Ten et al., 2007) showed that extra cytoplasmic abnormalities of the oocyte did not affect fertilization, embryo quality and clinical outcomes. According to the Istanbul consensus workshop on embryo assessment (Alpha Scientists in Reproductive Medicine and ESHRE Special Interest Group of Embryology, 2011) extra cytoplasmic anomalies (PBI morphology, PVS size, the appearance of the ZP) are simply phenotypic variations often related to in vitro culture and/or oocyte aging.

77

Studies by (Xia, 1997; Ebner et al. 1999; De Santis et al. 2005; Rienzi et al. 2008) however, suggested that the presence of extra cytoplasmic abnormalities of oocyte at MII stage (large and degenerated polar body in addition to large perivitelline space) are associated with a decreased potential of the cell to fertilize, cleave, and/or develop into a viable embryo. It has been suggested that a degenerated IPB may reflect an asynchrony between nuclear and cytoplasmic maturation which would explain the reduced ability of the cell to support pronuclear formation after ICSI. It has been postulated that the emission of an abnormally large IPB is due to the dislocation of the meiotic spindle. Large PVS may be ascribed to an over maturity of these oocytes at the time of ICSI (Rienzi et al. 2008; Cota et al., 2012). In this study implantation rate was statistically significant which was higher in the group with normal oocytes morphology 11.11% than the group of oocytes with extra cytoplasmic abnormalities 2.38%. But, in studies by (Balaban et al., 1998; Chamayou et al. 2006; Ten et al., 2007) demonstrated that extra cytoplasmic abnormalities had no effect on implantation and clinical pregnancy rates.

78

4.2 Conclusion:  Controlled ovarian stimulation protocols produce oocytes of great heterogeneity in both number and quality.  The fertilization, cleavage and high quality embryos rates were not affected by these high percentages of abnormal oocytes morphology. 

Implantation rate was higher in the group with normal oocytes morphology than the group with abnormal oocyte morphology.

79

4.3 Recommendations: 1. The evaluation of oocyte morphology before ICSI is strongly recommended in all scoring systems applied in IVF laboratories. 2. Oocyte scoring system applied by dividing abnormal MII oocyte into 3 subgroups : I. II. III.

MII oocytes with cytoplasmic abnormalities. MII oocytes with extra cytoplasmic abnormalities. MII oocytes with combined abnormalities. And in each subgroup grading for the oocytes applies in order to select which grade of oocyte to proceeded to ICSI

3. Further studies with larger size samples and adequate time is needed. 4. Genetic and epigenetic studies is essential to achieve better results and conclusions.

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Appendix

Name of wife:

age:

Name of husband:

age:

Year of infertility:

infertility factor:

Protocol:

trial:

Sperm source………………Preparation………………..Volume……………. Liquefaction…………………viscosity…………………PH…………… ….. Count…………………………Motility: A………..B……..C…….D………. Morphology: normal………………..Abnormal: head………..tail………… Middpeice…….. Hormonal assay: FSH: ….. LH: ..… P4:…. E2: …..E2 on day of HCG injection…………… Duration of induction…………………No. of oocytes retrieved………….. Classification of oocytes retrieved…….. MII………..

MI…………..GV………….

MII oocytes assessment…………. No. of normal MII

injected

fertilized

cleaved

Grade of embryos

transferred

No. of abnormal MII

Injected

fertilized

No. of abnormal MII with cytoplasmic abnormalities

Injected

No. of abnormal MII with extra cytoplasmic abnormalities

Injected

cleaved

fertilized

cleaved

fertilized

cleaved

Grade of embryos

Grade of embryos

Grade of embryos

No. of embryos transferred…………… BHCG…………………..Ultrasound: GS…………………

transferred

transferred

transferred

‫الخالصة‬ ‫الهدف من الدراسة‪ :‬هي تقييم تأثيرات شكل البويضة الناضجة و مدى تأثيرها على األصخاا و‬ ‫تكوين األجنة و نسبة غرس األجنة في الرحم من صخالل عملية الحقن المجهري‬ ‫طريقة العمل‪:‬‬ ‫تم تجميع البويضات من ‪ 231‬حالة صخاصة بأناث قد تم ءاجراء عملية الحقن المجهري لهن‬ ‫واللواتى يعانون من العقم‪ .‬وقد تم األستثناء عن االناث اللواتى ازواجهن ماابين بعقم رجالي‪ .‬و‬ ‫من مجموع ‪ 2111‬بويضة تم تجميعها ‪ 2101‬منها كانت بويضة ناضجة و التى تمت تقيمها‬ ‫للدراسة‪ .‬تم تقسيم هذه البويضات الناضجة حسب اشكالها الى‪:‬‬ ‫‪.2‬البويضة الناضجة ذات الشكل الطبيعي و التي تتميز بسايتوبالزم متجانس ولها صخلية قطبية‬ ‫واحدة‪.‬‬ ‫‪ .1‬البويضة الناضجة ذات الشكل الغيرالطبيعي للسايتوبالزم البويضة التي تشمل‪ (:‬سايتوبالزم‬ ‫غامق و ذو حبيبات و وجود جسم ضمين و تجاويف داصخل السايتوبالزم )‬ ‫‪ .3‬البويضة الناضجة ذات الشكل الغيرالطبيعي لخارج السايتوبالزم و التي تشمل‪ (:‬الشكل‬ ‫الغيرالطبيعي للخلية القطبية األولى و الشكل الغيرالطبيعي للمحيط الخارجي للسايتوبالزم)‬ ‫‪ .4‬البويضة الناضجة و التي تضم الشكل الغير الطبيعي للسايتوبالزم وصخارجها‪.‬‬ ‫النتيجة‪:‬‬ ‫و قد تبين من النتائج أن معدل عمر األناث كانت (‪ )0.032±33.33‬وانه من مجموع ‪2101‬‬ ‫بويضة ناضجة قد تم تقيميها ‪ )%20.14(231‬لديها شكل طبيعي بينما ‪ )%31.30(301‬بويضة‬ ‫لديها شكل غير طبيعي‪ .‬الشكل الغيرالطبيعي للسايتوبالزم البويضة الناضجة وجدت في‬ ‫‪ )%03.3(021‬بويضة أما الشكل الغيرالطبيعي لخارج السايتوبالزم فوجدت في‬ ‫‪ )%22.30(214‬أما البقية كانت للبويضات اللواتي تضم كال الشكلين الغير الطبيعيين‬ ‫للسايتوبالزم و صخارجها ‪)%13.11( 101‬‬ ‫النتائج لم تبين اي فرق هام في األصخاا وأنقسام الخاليا وتكوين األجنة منها اجنة ذات جودة‬ ‫عالية بين البويضات ذات الشكل الطبيعي و الغير الطبيعي و انما هناك فرق هام في نسبة غرس‬ ‫األجنة في الرحم بين المجموعات‪.‬‬ ‫األستنتاج‪:‬‬ ‫قد استنتج من هذه الدراسة ان كال من األصخاا وأنقسام الخاليا وتكوين األجنة لم تتأثر بشكل‬ ‫البويضة وانما نسبة غرس األجنة في الرحم زادت لدى األناث اللواتي شكل بويضهن طبيعية‪.‬‬

‫تة‬ ‫ئامانجى تويذينةوة ‪:‬‬ ‫هةلسةنطاندنى كاريطةرى شيوةى هيلكةى كامل لةسةر ثيتاندنى تاقيطةى و دةرئةنجامةكةى‬ ‫ريطاى كار‪:‬‬ ‫هةلساين بة وةرطرتنى هيلكةكان لة( ‪ ) 132‬ئافرةت كة ثيتاندنى تاقيطةيان ب كراوة بة مةرجي‬ ‫هاوسةرةكانيان نةز كيان نةبووبيت لة ك ى ( ‪ )1200‬هيلكةى ك كراوة ( ‪ )1056‬دانة هيلكةى كامل بوو كة‬ ‫هةلسةنطاندنيان ب كراوة وة ئةم هيلكة كامالنة دابةشكراون بة طويرةى شيوةكانيان ب ‪:‬‬ ‫‪)1‬‬ ‫‪)2‬‬

‫‪)3‬‬ ‫‪)4‬‬

‫شيوةى ئاسايى هيلكةى كامل‬ ‫شيوةى نائاسايى سايت ثالزمى هيلكةكة كة ئةمي دابة دةبيت ب‬ ‫أ‪ -‬سايت ثالزمى دةنك لةدار‬ ‫ بوونى تةنى دةنك لةلةى و ب شايى لة ناو سايت ثالزمدا‪..‬‬‫شيوةى ناسايى ديوى دةرةوةى سايت ثالزم كة ئةوي ثي ديت لة ‪:‬‬ ‫صخانةى جةمسةرى و ب شايى دةورى سايت ثالزم‪.‬‬ ‫ئةو هيلكة كامالنةى كة شيوةى نا ئاساييان هةية لة ديوى ناوةوةو دةرةوةى سايت ثالزمدا‬

‫ة جا‬ ‫لة ئةنجامدا ب مان دةركةوت كة ريذةى طشتى تةمةنى ئافرةتةكان (‪.)5.581 ±33.98‬‬ ‫هةروةها ب مان دةركةوت لة ك ى( ‪ )1056‬هيلكةى كامل ‪ )%17.04 ( 180‬هيلكة شيوةيان ئاسايى بووة ‪876‬‬ ‫(‪ )%82.95‬هيلكة شيوةى نائاسايى بووة لة م هيلكة شيوة نائاسايية ‪ )%58.9( 516‬هيلكة سايت ثالزميان‬ ‫ئاسايى نةبووة و ‪ )%11.87( 104‬دانة ديوى دةرةوةى سايت ثالزميان نائاسايى بووة وة ئةو ريذةية كة‬ ‫ماوةتةوة كة دةكاتة ‪ )%29.22 ( 256‬لة هةردوو ج رةكةى تيدا بووة (واتة ديووى دةرةوة و ناوةوةى‬ ‫سايت ثالزميان نائاسايى بووة ) بةم شيوةية ب مان دةردةكةويت كة هي جياوازيةكى بةرضاو نية لة نيوان‬ ‫ثيتاندن و دابة بوونى صخانةكان و دروست بوونى ك رثةلة لةطةل هيلكةى ئاسايى و نائاسايدا بةالم‬ ‫جياوازيةكى بةرضاو هةية لة ريذةى ضاندنى ك رثةلة لةناو رةحمى دايكةكةدا لةطةل هيلكةى ئاسايى و‬ ‫نائاسايدا‬ ‫ر ة جا ‪:‬‬ ‫لةو دةرئةنجامةوة ب مان روون بوةوة كة شيوةى هيلكةى كامل هي كاريطةريةكى نية لةسةر ثيتاندن و‬ ‫دروست بوونى ك رثةلة (ك رثةلةى كواليتى با ) بةالم كاريطةرى هةية لةسةر ريذةى ضاندنى ك رثةلة‪.‬‬

‫وةزارةتى صخويندنى باال و تويذينةوةى زانستى‬ ‫زانك ى سليمانى ‪ /‬فاكةلتى زانستة ثزيشكيةكان‬ ‫سكولى ثزيشكى ‪ /‬بةشى تويكارى‬

‫لي لينة‬

‫لةسةر ةي‬ ‫ر‬

‫تويذينةوةكة ثيشكة‬

‫د‬

‫ي‬

‫ي ا‬

‫ا‬

‫ر ةلة ري‬

‫ي ة لةطة يتا د‬

‫ا د لة ا‬

‫يتا د‬

‫اة‬

‫ا ي ةي‬

‫كراوة بة سكولى ثزيشكى زانك ى سليمانى وةك بةشي لة ثيداويستى بةدةست‬ ‫هينانى بروانامةى ماستةر لة زانستى ك رثةلةزانيدا‬

‫لة ية‬ ‫تان ئازاد سال‬ ‫‪M.B.Ch.B‬‬ ‫‪2005‬‬ ‫سةر ةر ت‬

‫را‬

‫لة ية‬

‫د‪.‬عيماد غانم قاسم‬ ‫ى‪.‬ثر فيس رى تويكارى‬

‫كةالويز‬

‫رمضان‬

‫تموز‬

‫‪2417‬‬

‫‪1435‬‬

‫‪2014‬‬