Growth and Differentiation of Small Ovarian Follicles in Mammals: Problems and Future Perspectives

Journal of Reproduction and Development, Vol. 48, No. 5, 2002 Review Growth and Differentiation of Small Ovarian Follicles in Mammals: Problems and Future Perspectives Sandra CECCONI 1) 1) Department of Biomedical Sciences and Technologies, University of L Aquila, 67100 L Aquila, Italy Abstract. The in vitro development of mammalian ovarian follicles still represents a challenge. Results achieved in the past several years have clarified some important aspects concerning the regulation of follicle and oocyte development, but the complexity of the process leading in vivo to the production of a fertilizable egg makes quite difficult to obtain large numbers of fully-grown mature germ cells starting from in vitro cultured follicles. The main problem is that culture systems currently utilized do not assure satisfactorily a co-ordinate development of both the somatic and germinal components of the ovarian follicle. An ideal culture media might be specially designed by the addition of specific components in correct ratio, in the attempt to reflect the dynamic changes which characterize follicle development. This is important, especially in the light of the fact that the period of culture could vary considerably, depending on the timing of follicle growth in vivo in the various species, being also influenced by the age of donors, methods and aims of culture. Despite this, in vitro technology represents not only an important tool to understand regulative processes underlying follicle development, but also a future option for the preservation of fertility. The present paper reviews current knowledge on advances and problems relevant to the culture of primordial and preantral mammalian follicles. Key words: Follicle, Oocyte, In vitro culture, Cryopreservation (J. Reprod. Dev. 48: 431 445, 2002) An Overview of Oocyte and Follicle Development The principal aim of oogenesis is the production of a fertilizable oocyte. In the mammalian ovary, more than 99% of oocytes undergo a degenerative process known as atresia [1], and in primates usually one preovulatory follicles is produced at every single ovarian cycle [2]. In order to establish a viable pregnancy, the oocyte needs to develop three fundamental properties, represented by the capacity of resuming and completing meiosis up to metaphase 2 (i.e nuclear maturation), undergoing Accepted for publication: July 12, 2002 fertilization and embryogenesis (i.e. cytoplasmic maturation), and acquiring a correct genomic imprinting (i.e. genomic maturation). Germ cells enter meiosis during fetal life, but remain arrested at the prophase of the first meiotic division, the germinal vesicle (GV) stage. Depending on the species, GV arrest may last from weeks to years [2]. Meiotic competence is progressively acquired during oocyte growth [3, 4], essentially as an oocyte autonomous property [4, 5]. In particular, the ability to spontaneously resume meiosis is acquired as the oocyte approaches its final size [6, 7], a stage which in the mouse usually coincides with antrum formation. Meiotic

432 CECCONI maturation is a two-step process, as the oocyte firstly resumes the meiotic process, evidenced by germinal vesicle breakdown (GVBD), and then completes this process arresting at metaphase II (MII) [6 8]. However, the observation that, in vitro, only a proportion of MII-arrested oocytes derived from small antral follicles are competent to be fertilized and develop up to blastocyst stage [9 12] indicates that completion of nuclear maturation is not sufficient to assure normal fertilization and embryonic development [10, 13]. The term cytoplasmic maturation includes the occurrence of stage-specific processes, such as the synthesis of specific proteins [14], the ability to release cortical granules [15], to release calcium from intracellular stores [16], to re-localize mitochondria [17] and to decondense sperm head [13, 18], which significantly contribute to successful fertilization. Similar to what occurs to the nucleus, maturation of the cytoplasm is acquired in a step-wise manner [19], requiring complete antral development [11] and maintenance of functional gap junctions between somatic and germ cells [10, 11, 20]. Also the release of GV content within the cytoplasm may have a role in determining the fate of fertilized eggs, and the compartmentalization of regulatory proteins into the nucleus or the cytoplasm may play an important role in determining egg polarity and subsequent embryo development [21 23]. As above mentioned, oocyte growth and differentiation rely upon the physiological development of granulosa cells. The existence of a bi-directional dialogue between germ and somatic cells is a sine qua non condition to obtain fertilizable oocytes [20, 24, 25]. Such a cross-talk utilizes two main systems of interactions, based on the presence of gap junctions (Fig. 1 [26]) as well as the production of paracrine and autocrine factors [Table 1; 24, 25, 27, 28]. Gap junctions are channels composed of connexins (Cx), a family of integral membrane proteins that mediate most of the granulosa cell-to-oocyte communications [29, 30]. The role of two classes of Cx, Cx43 and Cx37, has been extensively studied by using knockout mice. The disruption of the Gja1 gene encoding Cx43, Fig. 1. Mouse secondary follicle. Follicle cell (F) projections are send towards the oolemma (O). 700 (Reproduced with permission of the authors from G. Macchiarelli et al., 1992). Table 1. Bi-directional communications between oocyte and granulosa cells control functions necessary for successful development of both cell types GRANULOSA CELL FUNCTIONS REGULATED BY THE OOCYTE * Follicle organization * Proliferation, differentiation, growth-promoting activity * Steroidogenesis * Cumulus expansion, prostaglandin production OOCYTE FUNCTIONS REGULATED BY GRANULOSA CELLS * Growth * Meiotic arrest * Maturation * Transcriptional activity * Protein synthesis and phosphorylation

REVIEW: SMALL OVARIAN FOLLICLES IN VITRO 433 which is contained in the junctions between granulosa cells, causes neonatal lethality [31] and arrest of folliculogenesis at an early stage of development [32], due to the very low number of germ cells present in the ovary [33]. The targeted deletion of the Gja4 gene encoding Cx37 [34], which characterizes channels between the oocyte and granulosa cells, causes arrest of follicle development at the secondary stage, while oocytes are prevented from growing beyond about 74% of their final size [35]. As a consequence, the ovaries of these mice lack mature follicles, showing premature luteinization [34]. By means of heterologous gap junctions, granulosa cells support germ cell growth [20, 24], prevent premature meiotic resumption by transferring cell cyclearresting factors, as purines and camp [36, 37], modulate oocyte transcriptional activity [38], protein synthesis and phosphorylation [39 41], thus promoting oocyte capacity to undergo successful fertilization and preimplantation development. In humans, granulosa lutein cells influence oocyte quality and embryo developmental potential [42 44]. At the same time, the oocyte regulates a broad range of granulosa cell functions through the release of paracrine signals. Germ cells stimulate granulosa cell proliferation and differentiation [45 48], modulates steroidogenesis [49], expression of the c-kit ligand [50], LH receptor (LHR [51]), hyaluronic acid (HA) synthesis [52] and urokinasetype plasminogen activator (upa [53]). As antral follicle is formed, a gradient of oocyte-secreted factors directs the specialization of the surrounding granulosa cells, referred to as cumulus cells, in a specific phenotype, in order to create a microenvironment capable to better respond to the oocyte requirements. In particular, the oocyte is responsible for a low expression of LHR and upa production in cumulus cells, while enhancing their proliferative activity and HA secretion, and regulating prostaglandin-endoperoxide synthase 2 [54] as compared with mural granulosa cells. An increasing body of evidence suggests that the action of some oocyte-derived regulative factors is temporally restricted, depending on the developmental phase of either the somatic or the germinal compartments. This conclusion is supported by recent findings obtained in the mouse showing that only fully grown immature oocytes are capable of modulating matrix deposition/ degratation [53], and of secreting a specific granulosa cell mitogen [48]. This last observation is in contrast with previous results by Vanderhyden and collaborators [45], who reported that growing oocytes are capable of promoting granulosa cell proliferation and differentiation. Moreover, fullygrown oocyte-released factor(s) are capable of suppressing preantral granulosa cell ability to sustain the growth of growing oocytes, even though the maximum impairment of oocyte growth requires functional gap junctions [55]. Recently, Eppig and collaborators [56] have demonstrated that growing mouse oocytes isolated from preantral follicles are capable of inducing a significant increase in the rate of primordial follicles development. This results strongly reinforces the idea that germ cells play a leading role in the regulation of ovarian follicular development. To date, growth differentiation factor-9 (GDF-9 [57 63]), which belongs to the transforming factor (TGF β) family, is the first oocyte-derived factor identified with a well-established influence on mammalian female fertility. An other member of this family is bone morphogenetic protein 15 (BMP- 15), which mrna is expressed in sheep oocyte and regulates early folliculogenesis [64 66]. In rodents, GDF-9 mimics oocyte ability to induce cumulus expansion, hyaluronan synthase 2, cyclooxygenase 2 and steroidogenic acute regulator (StAR) mrnas, and to suppress upa and LHR-mRNA synthesis [59, 61]. In vivo treatment of GDF-9 null rats with recombinant GDF-9 (rgdf-9) results in significant stimulation of follicular growth [62]. The oocyte plays a role also in the process of ovulation by regulating progesterone production through the sequential release of an anti-luteinization factor and GDF-9, which suppresses [49, 51] or stimulates [59] steroid release, respectively. Finally, the acquisition of oocyte developmental competence requires also the occurrence of specific modifications necessary to imprint a small number of genes which have a critical role in the regulation of embryonic growth [67]. In mouse oocytes, imprinting occurs during growth phase [68], spanning from the primary to the antral stage [69]. Defects of correct imprinting leads to specific perturbations of gene expressions which disrupt establishment of allele-specific epigenetic modifications and dramatically affect embryonic outcome [70].

434 CECCONI Growth of Small Ovarian Follicles In vivo In the neonatal mammalian ovary primordial follicles are present in large numbers, and constitute the source of oocytes that have potential to ovulate during female reproductive life. They are formed by a quiescent oocyte arrested at the GV stage and surrounded by a single layer of flattened pre-granulosa cells. In many species, including humans, about 50 70% of oocytes die before follicle formation, likely as a consequence of major chromosomal abnormalities or defective mitochondria [71, 72]. Little is known about the cell-cell signalling molecules necessary for the organization of primordial follicle units and for their recruitment from the resting to the growth phase. The fact that growth to primary stage can occur without the action of extragonadal factors [73, 74] indicates that the local production of several interacting follicle/ oocyte-derived factors is likely to play a key role in the control of this process. The FIG α gene, expressed by the oocyte, controls follicle development and the production of zona pellucida proteins [75]. Likewise, Müllerian inhibiting substance (MIS [76]), basic fibroblast growth factor (bfgf [77]) and kit ligand (KL [78 82]) are involved in triggering primordial follicle development. As above mentioned, also GDF-9 plays a fundamental role since early stages of follicle development, because this factor is released by oocytes beginning from stage 3a, and its disruption causes a block in follicular development at stage 3b [57, 60, 83]. However, whether GDF-9 controls or not the transition from primordial to primary stage is still controversial, since GDF-9 mrna and protein are absent in primordial follicles of humans [83], but not in bovine and ovine follicles [84]. Indeed, in rodents primary follicles GDF-9 acts at the same time as modulator of theca cells differentiation [62] and of KL secretion [59]. This conclusion is sustained by GDF-9 capacity to stimulate the appearance of theca marker CYP17, and by the presence of oocyte larger than control in GDF-9 deficient rat due to up-regulation of KL. Treatment with rgdf-9 of deficient mice induces the progression from primordial to small preantral stage [62], a stage in which this paracrine factor collaborates with FSH in promoting further follicle differentiation and survival. It is generally accepted that early phases of folliculogenesis are independent from FSH, as small follicles grow even in hypophysectomized and hypogonadotrophic animals [85, 86]. Furthermore, FSH receptors (FSHR) are undetectable in primordial follicles, whilst they are recognizable in primary follicles and their expression increases significantly during the transition to the preantral stage [87 91]. Two alternative signalling pathways have been proposed to mediate the acquisition of FSHR [90]. The former, based on a camp-independent mechanism, is believed to be mediated by activin, a member of the TGF β family, which induces the expression of FSHR in cultured granulosa cells. The latter is thought to operate directly via camp, because this messenger is capable of stimulating aromatase synthesis and FSHR expression in pregranulosa cells and primordial follicles, as well as the production of the growth factor KL. The neurotransmitters vaso-intestinal peptide (VIP) and nor-epinephrine (NE), that act via a camp generating system [92], may also participate in early follicular development. Indeed, the first follicles to start growing are located in the more densely innervated ovarian region [1], and neonatal sympathectomy results in follicle arrest, reduced steroidogenesis and delayed puberty [93]. Thus the possibility exists that follicles more directly exposed to these camp-dependent signals specifically activated by neurotransmitters may acquire FSHR and gonadotropin responsiveness more rapidly than the others. Although gonadotropins do not influence initiation of follicle growth, they are critical for complete follicle development [88, 91]. FSH and LH have complementary effects, as LH stimulates the production of androgens by theca cells, which are aromatised under FSH action in granulosa cells (two-cell theory). However while the role of LH is limited to later stages of folliculogenesis, that of FSH is exerted more widely throughout the entire process. In humans transcription of FSHR genes has been evidenced in 33% of primary and 2- layered follicles [94]. In mice, FSHR are expressed by day 7 of postnatal life, and FSH levels increase in coincidence with granulosa cell proliferation [91]. The use of mutant mice for FSHR genes confirms that low gonadotropin levels determine a significant reduction in the number of developing follicles and increase atresia of the remaining ones

REVIEW: SMALL OVARIAN FOLLICLES IN VITRO 435 Fig. 2. In vitro growth of manually-isolated mouse follicles cultured for 6-days in non-spherical system. A. day 0: freshly-isolated follicle with some stromal cells attached; B. day 6: antral-like cavity and cumulus cell-oocyte complex are easily recognizable. Bar = 100 µm. [91]. Moreover, FSH regulates the expression of cyclin D2, a G1 phase protein, with the consequence that mice lacking this gene are infertile and show follicles arrested at early preantral stage [95]. Although FSH stimulates also KL expression in preantral granulosa cells, oocyte growth occurs in the absence of this gonadotropin [28] as well as in the presence of antibodies to KL [27, 81]. The fact that KL is expressed in preantral follicles may support its role as a factor promoting oocyte survival and cytoplasmic maturity [96]. This is a very important function, since during this phase the oocyte accumulates specific resources for subsequent maturation, fertilization and embryonic development. In vitro The aim of in vitro follicle culture is to mimic the process occurring in vivo, in order to investigate the underlying physiology and generate a fully grown oocyte able to undergo successful maturation and fertilization. The first attempts to culture in vitro mammalian follicles have been performed by recovering follicles from enzymatically-digested mouse ovaries. These follicles, presenting few theca cells and damaged basement membrane, were cultured as oocyte-granulosa cell complexes (OGCCs), embedded into collagen or agar gels or placed onto collagen-treated porous membrane to maintain close association between the oocyte and granulosa cells [97, 98]. Significant results were initially reported by Eppig and Schroeder [18], who obtained live offspring after in vitro maturation (IVM), fertilization (IVF) and transfer of oocytes obtained from granulosa cell complexes isolated from 12-day-old mice. Alternatively, mouse preantral follicles can be mechanically dissected from the ovary [99 104]. Although a low number of follicles is recovered, the presence of an intact theca allows maintainance of a physiological microtopograph in the absence of supporting matrix. At the same time, the presence of stromal cells around the follicle improves its overall development. During culture, follicles grow at a rate similar to that in vivo, releasing high levels of estrogen, and following FSH stimulation form antral cavities (Fig. 2). Moreover, a high proportion of them can respond to LH [102] or to EGF [104] administration by mimicking ovulation, evidenced by the expansion of cumulus-corona cells and maturation of the oocyte. With the application of this culture method, the birth of live youngs after fertilization and embryo transfer has been reported [102]. Fertilizable oocytes can be obtained also by culturing intact follicles on flat tissue culture dishes [101]. In this case, adhering theca cells function as substrate for subsequent granulosa cells proliferation and outgrow from the basament membrane. Under this experimental

436 CECCONI condition, the loss of follicle spherical aspect does not interfere with antral cavity formation and the acquisition of full oocyte developemental potential. This method might improve access of nutrients and growth factors to the different follicular compartments in comparison with procedures involving whole follicle culture. In vitro culture of preantral follicles from large mammals has been performed by mean of the same techniques described for rodents. Preantral bovine follicles have been firstly isolated by enzymatic [105] or combined enzymatic/mechanical procedures [106], but these treatments adversely affect further follicle development, as demonstrated by the massive degeneration of oocytes at the end of culture period. By contrast, the absence of theca layer does not impair porcine follicle development, as antral cavity is formed after culture in collagen gel, accompanied by the production of oocytes capable of maturing up to MII stage, but unable of undergoing fertilization [107]. Mechanical dissection allows to obtain undamaged follicles, but in large animals it is a very time-consuming procedure, due to the difficulty of recovering a sufficient number of preantral follicles from the fibrous connective tissue of ovaries of these species. Preantral follicle development in vitro up to antral stages has been obtained for bovine [108], and sheep [109], even though meiotic competence of IVG oocytes results very limited. A high percentage (51%) of porcine oocytes obtained from IVG large preantral/early antral follicles can complete meiotic maturation to MII stage, 43% of them undergoing fertilization and 13% of fertilized oocytes progressing throughout embryonic development [110]. The ultrastructure and viability of mechanically isolated small preantral and large preantral/early antral bovine follicles reveals that more than 90% of follicles have degenerated oocytes by the end of culture, as evidenced by the presence of numerous vacuoles, lipid droplets, swollen mitochondria in the cytoplasm, and clustering of chromatin material [111]. Concerning humans, Roy and Treacy in 1993 reported the first results relating to development of enzymatically-isolated preantral follicles cultured in agar gels [112], a part of which formed antrallike cavities following FSH stimulation. Higher percentages of antrum formation (about 71%) have been obtained from manually isolated preantral and early antral follicles [113]. Fig. 3. Overview of follicle development. In this section of human ovarian cortex, an early developing follicle is surrounded by a nest of primordial follicles ( 450). (Reproduced with the permission of the authors from Motta et al., 2002) As primordial follicles are the most abundant population in the ovary (Fig. 3 [114]), great attention has been devoted to the possibility of culturing them, especially after experiments demonstrating their capacity to restore fertility following transplantation [115, 116]. Since it is impossible to sustain cultures of isolated primordial follicles, in vitro organ culture or cortical slices culture have been used to assist the transition from primordial to primary stage. A two-step system has been developed in which primordial follicles from mouse neonatal ovaries are firstly activated in organ culture to assure the transition to early preantral stage, and the enzymatically isolated OGCCs are further cultured on collagentreated inserts [73]. These IVG oocytes have been matured and fertilized in vitro, and after embryo transfer in foster mothers one live pup has been produced. Unfortunately, application of this technology to large mammals has so far failed to produce the transition to preantral stage, despite initiation of primordial follicle growth [108, 117 120]. Recently, a two step culture has been attempted in which newborn mouse ovaries have been firstly allotransplanted, in order to obtain preantral follicles from the grafted tissue. After recovery and culture in vitro, many of these IVG oocytes showed the ability to undergo GVBD and progress to MII, in the proportion 60% and 45% respectively, despite morphological observations revealed a reduced number of gap junctions between granulosa cells and oocyte [118].

REVIEW: SMALL OVARIAN FOLLICLES IN VITRO 437 Experimental Conditions and Their Role in Determining Successful in vitro Culture Role of gonadotropins Follicle culture media utilized for in vitro culture can be sorted into two groups, depending on the presence or absence of serum, which can be replaced by BSA as an alternative protein source. Although the addition of FSH is considered not important in promoting primary follicle development to preantral stage [116, 117, 119], it is necessary to assure preantral follicle survival, antrum development and stimulation of steroidogenesis [120]. In sheep, this gonadotropin exerts a significant dose-dependent effect, with the highest doses tested improving overall culture outcome [109]. Recombinant FSH (rfsh) is effective in producing a high incidence of antrum formation and developmentally competent oocytes from IVG mouse preantral follicles [121]. Vitt and colleagues [122] have fractionated human rfsh and separately tested on mouse preantral follicle development the three isohormone fractions obtained, designated as least, mid and less acidic. Results have demonstrated that the least acidic isoform strongly stimulates growth, while the other two specifically stimulates estradiol release and antrum appearance in mouse preantral follicles. However, at least in mice, oocyte competence to undergo fertilization and implantation can be acquired also in the absence of gonadotropin stimulation, while it is decreased significantly in the absence of serum [123]. Measurable consequences derived from of the addition of LH require the presence of intact theca, and may have significant effect on increasing mouse follicular growth and differentiation, probably by mediating some yet unknown aspects of antral cavity formation. When added in a timely manner, LH induces ovulation [99, 102], even though EGF seems to be more efficient [104]. Role of other growth factors Various other substances added to culture media stimulate follicle growth and differentiation, and at present the list of growth factors and their reciprocal combinations is quite long [101]. DbcAMP, hypoxanthine, relaxin [124], recombinant GDF-9 [62], insulin [101, 120] have been added to improve mammalian follicle growth in vitro, while ascorbic acid is added to reduce granulosa cell apoptosis [110, 125, 126]. Other growth factors seem to exert a sort of species-specific effect. For example, KL is successful in supporting mouse [127] but not human [128] follicle development; IGFs (I and II) enhance granulosa cell responsiveness in human [129] and rodents [130], but not in bovine [131]. Furthermore, activin stimulates follicle survival in rodents but its role in humans is still controversial [101]; TGFβ1 stimulates glucose metabolism and potentiates FSH action in human follicle culture [132], while inhibits FSH effects in rat follicles [77]. By contrast, the activation of primordial follicles in culture of cortical strips is not improved by addition of gonadotropins, camp or EGF [133]. The ideal culture system The reduced proportion of oocytes developing to term following in vitro culture points to the conclusion that the current systems do not assure the coordinate development of both germinal and somatic components. The observation that follicle growth and differentiation can be maintained for several days, while oocyte quality decreases significantly, confirms that some important granulosa cell-oocyte signalling pathways are blocked or abnormally regulated. Premature death of the only one mouse produced after the complete oocyte in vitro development from primordial stage [123] further underlines these conclusions. In addition, it appears that the parameters currently utilized to evaluate the efficiency of culture conditions, such as maturation and fertilization rate, normal embryonic development and even survival to term, are probably inadequate to predict the well-being and normal life span of individuals born from oocytes completely developed in vitro. An ideal culture system would assure generation of essential endocrine and paracrine signals, produced at the appropriate time and at levels adequate to induce in germ cells the changes necessary for nuclear, cytoplasmic and genomic maturation. However, evaluation of the effects exerted by the combination of gonadotropins, serum and growth factors on the acquisition of oocyte developmental competence evidences some unexpected results. It has been observed that the number of blastocysts formed from IVG-IVM oocytes is significantly lowered when culture medium is supplemented with the combination of serum and FSH respect to serum alone [125], thus

438 CECCONI pointing to a negative role of FSH on oocyte developmental capacity. Indeed, oocytes grown in the absence of FSH assume a chromatin configuration typical of the oocytes obtained from preovulatory follicles of gonadotropin-primed mice [38]. Also, the addition of FSH and insulin impairs embryo formation, while these supplements alone are ineffective [134]. Since both hormones contribute significantly to granulosa cell development, it is possible that their combination abolishes the production of those oocyte-derived factors capable of controlling and maintaining the expression of correct somatic cell phenotype. Indeed, the combination of FSH and insulin triggers an inappropriate differentiation of granulosa cells by dramatically modifying protein synthesis patterns [135]. Because granulosa cells play an active role in the regulation of oocyte transcriptional activity [38], their non-physiological differentiation is highly deleterious for the acquisition of oocyte full competence. Several studies in recent years have emphasized that gene expression could be altered by in vitro conditions, and that abnormal epigenetic modifications characterize many types of human cancers [136]. Although genomic imprinting is established after embryo implantation [67, 69], epigenetic modifications of imprinted genes occurs in the oocyte throughout the period from primary to antral stage [68, 70]. Imprinted genes are vulnerable to epigenetic errors caused by nuclear manipulation, IVG-IVM, extended period of culture and the presence of additives [137, 138]. Recently, it has been observed that in vitro culture of sheep embryos in the presence of serum or somatic cell support can significantly influence foetal growth by inducing large off-spring syndrome (LOS) in about 25% of live youngs [139]. Considering these results, the question is whether possible embryonic anomalies can be overcome by identifying adverse epigenetic effects caused by culture conditions, or vice versa problems with genomic imprinting are presently un-resolvable (Table 2). Future Perspectives: in vitro Culture of Follicles for Therapeutic Treatments of Woman Infertility In the last years costant therapeutic advances have significantly increased the chances of clinical Table 2. Factors influencing in vitro development of mammalian small follicles Age of the donor Size classes of follicles at retrieval Techniques for isolation: enzymatic, mechanical or combination of both Culture conditions: organ or individual culture (intact spherical or non-spherical follicles), oxygen tension, duration, components in basal medium Lack of objective criteria for the determination of follicular quality during and at the end of culture Epigenetic modifications of imprinted genes remission in cancer patients, but in young women this is generally achieved at the cost of permanent loss of their fertility, due to collateral damage caused by anticancer treatments to the viability of germ cells [140]. Primarily, germ cell cryopreservation, applied as a preventive measure, could offer the possibility to preserve fertility in these patients as well as other women at risk of premature loss of their ovarian function Technically, a possible alternative is represented by the cryopreservation of fully grown oocytes. Although some pregnancies have been achieved from stored mature [141] and immature [142] oocytes, it is believed that the developmental capacity of cryopreserved oocytes could be compromised by the freezing/thawing treatment, making the success rate of this approach presently not comparable with other forms of intervention. Fertilizable mouse oocytes have been obtained from antral follicles after cryopreservation of whole ovaries [143], but unfortunately this procedure cannot be currently applied to human ovaries because of their much larger size. Thus, attention has been devoted to procedures by which mouse mature oocytes can be obtained after culture in vitro of small follicles isolated from frozen/thawed tissue [144]. Preantral follicles have been cryopreserved either before or after isolation from surrounding stromal cells, and then cultured in vitro up to ovulation, to obtain fertilizable oocytes. Ultrastructural evaluation after freezing reveals that 25% of mouse preantral follicles undergoes major structural damage, the surviving population showing intact oocyte-granulosa cells contacts [145]. In large mammals, Newton and cowokers [146] reported that the same percentage (about 20%) of DMSO-cryopreserved and freshly-collected

REVIEW: SMALL OVARIAN FOLLICLES IN VITRO 439 sheep granulosa cell-oocyte complexes develops antra after 30 days of in vitro culture. Histological analysis performed soon after thawing, reveals that about 50% of DMSO-cryopreserved sheep preantral follicles is severely damaged in both the somatic and germinal components (Fig. 4), while almost all the fresh follicles show a well preserved morphology (not shown; Cecconi S, Barboni B, Berardinelli P; manuscript in preparation). In view of the limited success derived from the cryopreservation of isolated follicles, the storage of ovarian cortical tissue may represent a promising source of primordial and primary follicles [144, 147]. Recently, Wang and collaborators [148] described the successful transplantation in oophorectomized recipients rats of fresh or frozenthawed intact ovaries, connected to their fallopian tubes and the upper segment of the uterus, and the establishment of normal pregnancy. In cancer patients, a serious limitation to ovarian transplantation is represented by the type of cancer [149] and by the possibility of causing unintentional spreading following transplantation of the stored material. This aspect is still controversial [150, 151]. Indeed, the alternative option is represented by in vitro culture of follicles isolated from frozen/thawed cortical tissue, since this may overcome the problem of metastatic cells, due to the lack of stroma and vascularization in the isolated tissue. However, the present inability to obtain good quality oocytes from IVG follicles could be overcome by future improvement of this in vitro technology, and the identification of new cryoprotectants, pre-treatment of the grafts with specific growth factors [152] or antioxidants [153] which will contribute to reduce damages caused by reperfusion and transplantation. Conclusions Fig. 4. Photomicrographs of sheep preantral follicles. A. freshly isolated; B C. cryopreserved in DMSO. In C is evident the loss of gap junctional communications and a damaged cytoplasm in the oocyte. Bar= 50 µm. In this review the main aspects of in vivo and in vitro folliculogenesis have been focused. Alteration of epigenetic informations derived by in vitro culture represents the most important complication which questions the applicability of this strategy. How culture can lead to such a major effect is still unclear and require further study. Despite the urgent request of methods for restoring fertility in young women affected by cancer, we should keep in mind that the regulative processes we are approaching are considerably complex and need to be accurately examined. Therefore, it appears that, although it is important to improve the grafting procedures, the risk of reimplantation of metastatic foci cannot be underestimated, and that this option should be offered only to accurately selected patients. We are confidential that the use of animal models will help us to understand the causes impairing oocyte and embryo viability, to

440 CECCONI compensate for the adverse effects of in vitro procedures and to validate technologies before transfer to humans. Acknowledgments Coticchio, Riccardo Talevi and Guido Macchiarelli for critical reading of the manuscript, Dr.Ena Colantoni for technical help, and my sons Lorenzo and Valerio for their patience. My work cited in this review was supported by the Italian Ministry of University and Scientific Research. I would thank Drs. Rita Canipari, Giovanni References 1. Hirshfield AN. Development of follicles in the mammalian ovary. Int Rev Cytol 1991; 124: 43-101. 2. Gougeon A. Dynamics of follicular growth in the human: a model from preliminary results. Hum Reprod 1986; 1: 81-87. 3. Sorensen RA, Wassarman PM. Relationship between growth and meiotic maturation of mouse oocytes. Dev Biol 1976; 50: 531-536. 4. Chesnel F, Wigglesworth K, Eppig JJ. Acquisition of meiotic competence by denuded mouse oocytes: participation of somatic-cell product(s) and camp. Dev Biol 1994; 161: 285-295. 5. Canipari A, Palombi F, Riminucci M, Mangia F. Early programming of maturation competence in mouse oogenesis. Dev Biol 1984; 102: 519-524. 6. Szybek K. In vitro maturation of oocytes from sexually immature mice. J Endocrinol 54: 527-528. 7. Wassarman PM, Josefowicz WJ. Oocyte development in the mouse: an ultrastructural comparison of oocytes isolated at various stages of growth and meiotic competence. J Morphol 1978; 156: 209-235. 8. Tsafriri A, Channing CP. Influences of follicular maturation and culture conditions on the meiosis of pig oocytes in vitro. J Reprod Fert 1975; 43: 149-152. 9. Eppig JJ, Schultz RM, O Brien M, Chesnel F. Relationship between the developmental programs controlling nuclear and cytoplasmic maturation of mouse oocytes. Dev Biol 1994; 164: 1-9. 10. Eppig JJ, O Brien M, Wigglesworth K. Mammalian oocyte growth and development in vitro. Mol Reprod Dev 1996; 44: 260-273. 11. Cecconi S, D Aurizio R, Colonna R. Role of antral follicle development and cumulus cells on in vitro fertilization of mouse oocytes. J Reprod Fertil 1996; 107: 207-214. 12. Moor RM, Lee C, Dai YF, Fulka J Jr. Antral follicles confer developmental competence on oocytes. Zygote 1996; 4: 289-293. 13. Thibault C. Are follicular maturation and oocyte maturation independent processes? J Reprod Fertil 51: 397-411. 14. Schultz RM, Wassarman PM. Specific changes in the pattern of protein synthesis during meiotic maturation of mammalian oocytes in vitro. Proc Natl Acad Sci USA 1976; 74: 538-541. 15. Abbott AL, Xu Z, Kopf GS, Ducibella T, Schultz RM. In vitro culture retards spontaneous activation of cell cycle progression and cortical granule exocytosis that normally occur in in vivo unfertilized mouse eggs. Biol Reprod 1998; 59: 1515-1521. 16. Carroll J, Jones KT, Whittingham DG. Ca 2+ release and the development of Ca 2+ release mechanisms during oocyte maturation: a prelude to fertilization. Rev Reprod 1996; 1: 137-143. 17. Van Blerkom J, Runner M. Mithochondrial reorganization during resumption of arrested meiosis in the mouse oocyte. Am J Anat 1984; 171: 335-355. 18. Usui N, Yanagimachi R. Behaviour of hamster sperm nuclei incorporated into eggs at varius stages of maturation, fertilization and early development. The appearance and disappearance of factors involved in sperm chromatin decondensation in egg cytoplasm. J Ultrastruct Res 1976; 57: 276-288. 19. Eppig JJ, Schroeder AC. Capacity of mouse oocytes from preantral follicles to undergo embryogenesis and development to live young after growth, maturation, and fertilization in vitro. Biol Reprod 1989; 41: 268-276. 20. Buccione R, Schroeder AC, Eppig JJ. Interactions between somatic cells and germ cells throughout mammalian oogenesis. Biol Reprod 1990; 43: 543-547. 21. Antczak M, Van Blerkom J. Oocyte influences on early development: the regulatory proteins leptin and STAT3 are polarized in mouse and human oocytes and differentially distributed within the cells of the preimplantation stage embryo. Mol Hum Repro 1997; 3: 1067-1086. 22. Pines J. Four-dimensional control of the cell cycle. Nat Cell Biol 1999; 1: E73-79. 23. Kemphues K. PARsing embryonic polarity. Cell 2000; 101: 345-348. 24. Cecconi S, Colonna R. Influence of granulosa cells and of different somatic cell types on mammalian

REVIEW: SMALL OVARIAN FOLLICLES IN VITRO 441 oocyte development in vitro. Zygote 1996; 4: 305-307. 25. Eppig JJ. Oocyte control of ovarian follicular development and function in mammals. Reproduction 2001; 122: 829-938. 26. Macchiarelli G, Vizza E, Nottola SA, Familiari G, Motta PM. Cellular and microvascular changes of the ovarian follicle during folliculogenesis: a scanning electron microscopic study. Arch Histol Cytol 1992; 55: 191-204. 27. Cecconi S, Rossi G, De Felici M, Colonna R. Mammalian oocyte growth in vitro is stimulated by soluble factor(s) produced by preantral granulosa cells and by Sertoli cells. Mol Reprod Dev 1996; 44: 540-546. 28. Eppig JJ, Chesnel F, Hirao Y, O Brien MJ, Pendola FL, Watanabe S, Wigglesworth K. Oocyte control of granulosa cell development: how and why. Hum Reprod 1997; 12: 127-132. 29. Goodenough DA, Goliger JA, Paul DL. Connexins, connexons, and intercellular communication. Ann Rev Biochem 1996; 65: 475-502. 30. Simon AM, Goodenough DA. Diverse functions of vertebrate gap junctions. Trends Cell Biol 1998; 8: 477-483. 31. Chen ZQ, Lefebvre D, Bai XH, Reaume A, Rossant J, Lye SJ. Identification of two regulatory elements within the promoter region of the mouse connexin 43 gene. J Biol Chem 1995; 270: 3863-3868. 32. Ackert CL, Gittens JE, O Brien MJ, Eppig JJ, Kidder GM. Intercellular communication via connexin 43 gap junctions is required for ovarian folliculogenesis in the mouse. Dev Biol 2001; 233: 258-270. 33. Juneja SC, Barr KJ, Enders GC, Kidder GM. Defects in the germ line and gonads of mice lacking connexin43. Biol Reprod 1999; 60: 1263-1270. 34. Simon AM, Goodenough DA, Li E, Paul DL. Female infertility in mice lacking connexin 37. Nature 1997; 385: 525-529. 35. Carabatsos MJ, Sellitto C, Goodenough DA, Albertini DF. Oocyte-granulosa cell heterologous gap junctions are required for the coordination of nuclear and cytoplasmic meiotic competence. Dev Biol 2000; 222: 167-179. 36. Downs SM, Daniel SA, Bornslaeger EA, Hoppe PC, Eppig JJ. Maintenance of meiotic arrest in mouse oocytes by purines: modulation of camp levels and camp phosphodiesterase activity. Gam Res 1989; 23: 323-334. 37. Downs SM, Coleman DL, Ward-Bailey PF, Eppig JJ. Hypoxanthine is the principal inhibitor of murine oocyte maturation in a low molecular weight fraction of porcine follicular fluid. Proc Natl Acad Sci USA 1985; 82: 454-458. 38. De La Fuente R, Eppig JJ. Transcriptional activity of the mouse oocyte genome: companion granulosa cells modulate transcription and chromatin remodeling. Dev Biol 2001; 229: 224-236. 39. Colonna R, Cecconi S, Tatone C, Mangia F, Buccione R. Somatic cell-oocyte interactions in mouse oogenesis: stage-specific regulation of mouse oocyte protein phosphorylation by granulosa cells. Dev Biol 1989; 133: 305-308. 40. Cecconi S, Tatone C, Buccione R, Mangia F, Colonna R. Granulosa cell-oocyte interactions: the phosphorylation of specific proteins in mouse oocytes at the germinal vesicle stage is dependent upon the differentiative state of companion somatic cells. J Exp Zool 1991; 258: 249-254. 41. Motlik J, Nagai T, Kikuchi K. Resumption of meiosis in pig oocytes cultured with cumulus and parietal granulosa cells: the effect of protein synthesis inhibition. J Exp Zool 1991; 259: 386-391. 42. Plachot M, Antoine JM, Alvarez S, Firmin C, Pfister A, Mande-Ibaum J, Junca AM, Salat-Baroux J. Granulosa cells improve human embryo development in vitro. Hum Reprod 1993; 8: 2133-2140. 43. Fabbri R, Porcu E, Marsella T, Primavera MR, Cecconi S, Nottola SA, Motta P M, Venturoli S, Flamini C. Human embryo development and pregnancies in an homologous granulosa cell coculture system. J Ass Reprod Gen 2000; 17: 1-12. 44. Cecconi S, Rossi G, Gualtieri R, Talevi R. Presence of a 31-kd protein band in human cumulus-corona radiata-conditioned media and pregnancy outcome. Fert Ster 2001; 75: 1-11 45. Vanderhyden BC, Telfer EE, Eppig JJ. Mouse oocytes promote proliferation of granulosa cells from preantral and antral follicles in vitro. Biol Reprod 1992; 46: 1196-1204. 46. Lanuza GM, Fischman ML, Baranao JL. Growth promoting activity of oocytes on granulosa cells is decreased upon meiotic maturation. Dev Biol 1998; 197: 129-139. 47. Li R, Norman RJ, Armstrong DT, Gilchrist RB. Oocyte-secreted factor(s) determine functional differences between bovine mural granulosa cells and cumulus cells. Biol Reprod 2000; 63: 839-845. 48. Gilchrist RB, Ritter LJ, Armstrong DT. Mouse oocyte mitogenic activity is developmentally coordinated throughout folliculogenesis and meiotic maturation. Dev Biol 2001; 240: 289-298. 49. Vanderhyden BC, Cohen JN, Morley P. Mouse oocytes regulate granulosa cell steroidogenesis. Endocrinology 1993; 133: 423-426. 50. Joyce IM, Pendola FL, Wigglesworth K, Eppig JJ. Oocyte regulation of kit ligand expression in mouse ovarian follicles. Dev Biol 1999; 214: 342-353. 51. Eppig JJ, Wigglesworth K, Pendola F, Hirao Y. Murine oocytes suppress expression of luteinizing hormone receptor messenger ribonucleic acid by granulosa cells. Biol Reprod 1997; 56: 976-984. 52. Salustri A, Yanagishita M, Hascall VC. Mouse

442 CECCONI oocytes regulate hyaluronic acid synthesis and mucification by FSH-stimulated cumulus cells. Dev Biol 1990; 138: 26-32. 53. Canipari R, Epifano O, Siracusa G, Salustri A. Mouse oocytes inhibit plasminogen activator production by ovarian cumulus and granulosa cells. Dev Biol 1995; 167: 371-378. 54. Joyce IM, Pendola FL, O Brien M, Eppig JJ. Regulation of prostaglandin-endoperoxide synthase 2 messenger ribonucleic acid expression in mouse granulosa cells during ovulation. Endocrinology 2001; 142: 3187-3197. 55. Cecconi S, Rossi G. Mouse antral oocytes regulate preantral granulosa cell ability to stimulate oocyte growth in vitro. Dev Biol 2001; 233: 186-191. 56. Eppig JJ, Wigglesworth K, Pendola FL. The mammalian oocyte orchestrates the rate of ovarian follicular development. Proc Natl Acad Sci USA 2002, 99: 2890-2894. 57. McGrath SA, Esquela AF, Lee SJ. Oocyte-specific expression of growth/differentiation factor-9. Mol Endocrinol 1995; 9: 131-136. 58. Nishimori K, Matzuk MM. Transgenic mice in the analysis of reproductive development and function. Rev Reprod 1996; 1: 203-212. 59. Elvin JA, Clark AT, Wang P, Wolfman NM, Matzuk MM. Paracrine actions of growth differentiation factor-9 in the mammalian ovary. Mol Endocrinol 1999; 13: 1018-1034. 60. Elvin JA, Yan C, Wang P, Nishimori K, Matzuk MM. Molecular characterization of the follicle defects in the growth differentiation factor 9- deficient ovary. Mol Endocrinol 1999; 13: 1035-1045. 61. Vitt UA, Hsueh AJ. Stage-dependent role of growth differentiation factor-9 in ovarian follicle development. Mol Cell Endocrinol 2002; 186: 21-31. 62. Vitt UA, McGee EA, Hayashi M, Hsueh AJ. In vivo treatment with GDF-9 stimulates primordial and primary follicle progression and theca cell marker CYP17 in ovaries of immature rats. Endocrinology 2000; 141: 3814-3820. 63. Dong J, Albertini DF, Nishimori K, Kumar TR, Lu N, Matzuk MM. Growth differentiation factor-9 is required during early ovarian folliculogenesis. Nature 1996; 383: 531-535. 64. Dube JL, Wang P, Elvin J, Lyons KM, Celeste AJ, Matzuk MM. The bone morphogenetic protein 15 gene is X-linked and expressed in oocytes. Mol Endocrinol 1998; 12: 1809-1817. 65. Otsuka F, Yao Z, Lee T, Yamamoto S, Erickson GF, Shimasaki S. Bone morphogenetic protein-15. Identification of target cells and biological functions. J Biol Chem 2000; 275: 3952-3958. 66. Galloway SM, McNatty KP, Cambridge LM, Laitinen MP, Juengel JL, Jokiranta TS, McLaren RJ, Luiro K, Dodds KG, Montgomery GW, Beattie AE, Davis GH, Ritvos O. Mutations in an oocytederived growth factor gene (BMP15) cause increased ovulation rate and infertility in a dosagesensitive manner. Nat Genet 2000; 25: 279-283. 67. Barlow DP. Gametic imprinting in mammals. Science 1995; 270: 1610-1613. 68. Bao S, Obata Y, Carroll J, Domeki I, Kono T. Epigenetic modifications necessary for normal development are established during oocyte growth in mice. Biol Reprod 2000; 62: 616-621. 69. Obata Y, Kaneko-Ishino T, Koide T, Takai Y, Ueda T, Domeki I, Shiroishi T, Ishino F, Kono T. Disruption of primary imprinting during oocyte growth leads to the modified expression of imprinted genes during embryogenesis. Development 1998; 125: 1553-1560. 70. Obata Y, Kono T. Maternal primary imprinting is established at a specific time for each gene throughout oocyte growth. J Biol Chem 2002; 277: 5285-5289. 71. Morita Y, Tilly JL. Oocyte apoptosis: like sand through an hourglass. Dev Biol 1999; 213: 1-17. 72. Pepling ME, Spradling AC. Mouse ovarian germ cell cysts undergo programmed breakdown to form primordial follicles. Dev Biol 2001; 234: 339-351. 73. Eppig JJ, O Brien MJ. Development in vitro of mouse oocytes from primordial follicles. Biol Reprod 1996; 54: 197-207. 74. Wandji SA, Srsen V, Voss AK, Eppig JJ, Fortune JE. Initiation in vitro of growth of bovine primordial follicles. Biol Reprod 1996; 55: 942-948. 75. Soyal SM, Amleh A, Dean J. FIGalpha, a germ cellspecific transcription factor required for ovarian follicle formation. Development 2000; 127: 4645-4654. 76. McGee EA, Smith R, Spears N, Nachtigal MW, Ingraham H, Hsueh AJ. Mullerian inhibitory substance induces growth of rat preantral ovarian follicles. Biol Reprod 2001; 64: 293-298. 77. Nilsson E, Parrott JA, Skinner MK. Basic fibroblast growth factor induces primordial follicle development and initiates folliculogenesis. Mol Cell End 2001; 175: 123-130. 78. Huang EJ, Manova K, Packer AI, Sanchez S, Bachvarova RF, Besmer P. The murine steel panda mutation affects kit ligand expression and growth of early ovarian follicles. Dev Biol 1993; 157: 100-109. 79. Parrott JA, Skinner MK. Kit-ligand/stem cell factor induces primordial follicle development and initiates folliculogenesis. Endocrinology 1999; 140: 4262-4271. 80. Manova K, Huang EJ, Angeles M, De Leon V, Sanchez S, Pronovost SM, Besmer P, Bachvarova RF. The expression pattern of the c-kit ligand in gonads of mice supports a role for the c-kit receptor in oocyte growth and in proliferation of spermatogonia. Dev Biol 1993; 157: 85-99. 81. Klinger FG, De Felici M. In vitro development of growing oocytes from fetal mouse oocytes: stage-

REVIEW: SMALL OVARIAN FOLLICLES IN VITRO 443 specific regulation by stem cell factor and granulosa cells. Dev Biol 2002; 244: 85-95. 82. Yoshida H, Takakura N, Kataoka H, Kunisada T, Okamura H, Nishikawa SI. Stepwise requirement of c-kit tyrosine kinase in mouse ovarian follicle development. Dev Biol 1997; 184: 122-137. 83. Aaltonen J, Laitinen MP, Vuojolainen K, Jaatinen R, Horelli-Kuitunen N, Seppa L, Louhio H, Tuuri T, Sjoberg J, Butzow R, Hovata O, Dale L, Ritvos O. Human growth differentiation factor 9 (GDF-9) and its novel homolog GDF-9B are expressed in oocytes during early folliculogenesis. J Clin Endocrinol Metab 1999; 84: 2744-2750. 84. Bodensteiner KJ, Clay CM, Moeller CL, Sawyer HR. Molecular cloning of the ovine Growth/ Differentiation factor-9 gene and expression of growth/differentiation factor-9 in ovine and bovine ovaries. Biol Reprod 1999; 60: 381-386. 85. Halpin DM, Charlton HM, Faddy MJ. Effects of gonadotrophin deficiency on follicular development in hypogonadal (hpg) mice. J Reprod Fertil 1986; 78: 119-125. 86. Hirshfield AN. Comparison of granulosa cell proliferation in small follicles of hypophysectomized, prepubertal, and mature rats. Biol Reprod 1985; 32: 979-987. 87. Smith SS, Ojeda SR. Neonatal release of gonadotropins is essential for development of ovarian follicle-stimulating hormone receptors. Biol Reprod 1986; 34: 219-227. 88. Hillier SG. Current concepts of the roles of follicle stimulating hormone and luteinizing hormone in folliculogenesis. Hum Reprod 1994; 9: 188-191. 89. Dunkel L, Tilly J, Shikone T, Nishimori K, Hsueh AWJ. Follicle stimulating hormone receptor expression in the rat ovary: increases during prepubertal development and regulation by opposing actions of transforming growth factor beta and alpha. Biol Reprod 1994; 50: 940-948. 90. Mayerhofer A, Dissen GA, Costa ME, Ojeda SR. A role for neurotransmitters in early follicular development: induction of functional folliclestimulating hormone receptors in newly formed follicles of the rat ovary. Endocrinology 1997; 138: 3320-3329. 91. McGee EA, Hsueh AJ. Initial and cyclic recruitment of ovarian follicles. Endocr Rev 2000; 21: 200-214. 92. Advis JP, Ahmed CE, Ojeda SR. Direct hypothalamic control of vasoactive intestinal peptide (VIP) levels in the developing rat ovary. Brain Res Bull 1989; 22: 605-610. 93. Lara HE, Hill DF, Katz KH, Ojeda SR. The gene encoding nerve growth factor is expressed in the immature rat ovary: effect of denervation and hormonal treatment. Endocrinology 1990; 126: 357-363. 94. Oktay K, Briggs D, Gosden RG. Ontogeny of follicle stimulating hormone receptor gene expression in isolated human ovarian follicles. J Clin Endocrinol Metab 1997; 82: 3748-3751. 95. Sicinski P, Donaher JL, Geng Y, Parker SB, Gardner H, Park MY, Robker RL, Richards JS, McGinnis LK, Biggers JD, Eppig JJ, Bronson RT, Elledge SJ, Weinberg RA. Cyclin D2 is an FSHresponsive gene involved in gonadal cell proliferation and oncogenesis. Nature 1996; 384: 470-474. 96. Reynaud K, Cortvrindt R, Smitz J, Driancourt MA. Effects of Kit Ligand and anti-kit antibody on growth of cultured mouse preantral follicles. Mol Reprod Dev 2000; 56: 483-494. 97. Torrance C, Telfer E, Gosden RG. Quantitative study of the development of isolated mouse preantral follicles in collagen gel culture. J Reprod Fertil 1989; 87: 367-374. 98. Eppig JJ, Telfer EE. Isolation and culture of oocytes. Methods Enzymol 1993; 225: 77-84. 99. Boland NI, Humpherson PG, Leese HJ, Gosden RG. Pattern of lactate production and steroidogenesis during growth and maturation of mouse ovarian follicles in vitro. Biol Reprod 1993; 48: 798-806. 100. Cortvrindt R, Smitz J, Van Steiteghen A. In vitro maturation, fertilization and embryo development of immature oocytes from early preantral follicles from prepubertal mice in a simplified culture system. Hum Reprod 1996; 11: 2656-2566. 101. Smitz J, Cortvrindt R. The earliest stages of folliculogenesis in vitro. Reproduction 2002; 123: 185-202. 102. Spears N, Boland NI, Murray AA, Gosden RG. Mouse oocytes derived from in vitro grown primary ovarian follicles are fertile. Hum Reprod 1994; 9: 527-532. 103. Spears N. In-vitro growth of ovarian oocytes. Hum Reprod 1994; 9: 969-970. 104. Boland NI, Gosden RG. Effects of epidermal growth factor on the growth and differentiation of cultured mouse ovarian follicles. J Reprod Fertil 1994; 101: 369-374. 105. Wandji SA, Eppig JJ, Fortune JE. FSH and growth factors affect the growth and endocrine functions in vitro of granulosa cells of bovine preantral follicles. Theriogenology 1996; 45: 817-832. 106. Figureido JR, Hulshof SCJ, Van den Hurk R, Ectors FJ, Fontes RS, Nusgens B, Bevers MM, Beckers JF. Development of a new mechanical and enzymatical method for the isolation of intact preantral follicles from fetal, calf and adult bovine ovaries. Theriogenology 1993; 40: 789-799 107. Hirao Y, Nagai T, Kubo M, Miyano T, Miyake M, Kato S. In vitro growth and maturation of pig oocytes. J Reprod Fertil 1994; 100: 333-339. 108. Telfer EE. The development of methods for