Abstract
Ovarian cryopreservation rapidly developed from basic science to clinical application and can now be used to preserve the fertility of girls and young women at high risk of sterility. Primordial follicles can be cryopreserved in ovarian cortex for long-term storage and subsequently autografted back at an orthotopic or heterotopic site to restore fertility. However, autografting carries a risk of re-introducing cancer cells in patients with blood-born leukaemias or cancers with a high risk of ovarian metastasis. For these women fertility restoration could only be safely achieved in the laboratory by the complete in vitro growth (IVG) and maturation (IVM) of cryopreserved primordial follicles to fertile metaphase II (MII) oocytes. Culture systems to support the development of human oocytes have provided greater insight into the process of human oocyte development as well as having potential applications within the field of fertility preservation. The technology required to culture human follicles is extremely challenging, but significant advances have been made using animal models and translation to human. This review will detail the progress that has been made in developing human in vitro growth systems and consider the steps required to progress this technology towards clinical application.
Introduction
It is now over 40 years since the birth of the first IVF baby (Niederberger et al. 2018) and since then several advances have been made in the area of assisted reproductive technologies particularly the emergence of the new field of fertility preservation (Anderson & Baird 2019). Women now have the opportunity to have pieces of their ovarian cortex removed and stored prior to damaging chemotherapy treatments with the goal of future re-implantation, however, for some patients re-implantation carries a risk of re-introducing malignant cells (Anderson et al. 2017). The development of an ‘artificial ovary’ system is ongoing in an attempt to avoid the risk of re-introducing malignant cells. The artificial ovary involves isolating follicles and reconstituting them with somatic cells within a matrix in the hope that any remaining malignant cells would be removed before transplantation (Dolmans & Amorim 2019). The ability to support oocyte development entirely in vitro would offer an alternative strategy by avoiding the need for transplantation to restore fertility. This review outlines the potential and difficulties of developing culture systems to support the development of human oocytes.
The ovarian tissue that is harvested for cryopreservation contains non-growing follicles at the primordial stage of development. These follicles contain the earliest stage of oocyte and research on how to develop them entirely in vitro has been ongoing for many years. Culture systems for in vitro gametogenesis/growth (IVG) need to support all stages of oocyte development from activation of dormant primordial follicles to a stage where oocytes can undergo meiotic maturation and be fertilised. If this methodology could be demonstrated to be safe it would maximise the potential of cryopreserved ovarian tissue and have many clinical applications relevant to fertility preservation and assisted reproduction. Culturing immature oocytes would negate the need for hormonal stimulation of the patient, avoid multiple surgical procedures to harvest oocytes but most importantly there would be options for fertility preservation/restoration to women who currently have none (Anderson et al. 2017, Telfer 2019).
The clinical application of growing primordial follicles in vitro would be especially relevant to prepubertal girls having ovarian tissue removed and cryopreserved prior to being exposed to potentially gonadotoxic treatments (Anderson et al. 2015, 2017). Freezing mature oocytes or embryos is not possible for prepubertal girls therefore their only option for fertility preservation is cryopreservation of ovarian cortical tissue with the potential for subsequent re-implantation (Anderson et al. 2015, 2017). If re-implantation was contra-indicated, IVG would provide an additional option for restoration of fertility in such cases. Given that cryopreserved cortical strips contain predominantly primordial follicles, it is the obvious starting material for a culture system; however, this also presents many challenges. The realisation of clinical applications of IVG will require a great deal of research to optimise culture systems and to demonstrate the safety of these techniques. Nonetheless progress in developing culture techniques to support in vitro development of human oocytes has been made.
Producing competent oocytes for assisted reproduction technologies (ARTs) is the main objective of IVG systems, but the development of these methods provides models of human gametogenesis that allow for detailed analyses of the sequential stages of human oocyte development of which we still know very little about. In addition, culture systems facilitate analysis and assessment of cryopreserved ovarian tissue prior to transplantation, making IVG central to fertility preservation programmes (Picton et al. 2008, Telfer & McLaughlin 2011).
Follicle/oocyte development
The most immature stage of oocytes are contained within structures known as primordial follicles (Fig. 1). In most mammals follicle formation occurs before birth and these form the store of female germ cells that will be utilised throughout reproductive life (Telfer & Zelinski 2013). Recruitment of follicles to growth is controlled by a defined sequence of events regulated by paracrine and endocrine factors (Hsueh et al. 2015). Initiation of follicle growth is also influenced by the intra ovarian environment and in particular biomechanical factors (Shah et al. 2018). The key stages that need to be supported can be summarised as (1) activation of primordial follicle growth, (2) development to multi-laminar pre-antral follicle stage, (3) antral cavity formation and differentiation of granulosa cells to mural and cumulus cells, (4) Proliferation of granulosa cells and expansion of the fluid filled antral cavity leading to the formation of the pre-ovulatory or Graafian follicle, and (5) release of the cumulus-oocyte complex after rupture of the Graafian follicle in response to the surge of luteinising hormone (LH) triggering ovulation (Fig. 1) (Telfer & Zelinski 2013).
Representation of stages of oocyte formation and development. Upper panel: proliferation of primordial germ cells (PGCs) which form nests and then enter meiosis to the dictyate stage of Prophase 1 forming oocytes associated with somatic cells to form primordial follicles. In humans this occurs during fetal life. Lower panel: Primordial follicles are continually activated to grow throughout reproductive life and progress through follicular development with co-ordinated oocyte development. Sizes shown illustrate oocyte development. Reproduced from Anderson & Telfer (2018) with permission.
Citation: Reproduction 158, 5; 10.1530/REP-19-0077
The majority of follicles in all mammals will be at the primordial stage and during follicle formation the oocyte is arrested at Prophase I of meiosis and remains in meiotic arrest until ovulation. During follicle growth and development the oocyte needs to acquire the competence to resume meiosis as well as the ability to support events during fertilisation and early embryonic development. This process is referred to as the acquisition of developmental competence and the development of culture conditions to support these processes represents an enormous technical challenge (Anderson & Telfer 2018) that requires a good understanding of the complex requirements of the various compartments within the ovary that is, oocyte, granulosa, theca and stromal cells. These requirements are dynamic and complex (Anderson & Telfer 2018). To ensure the development of healthy oocytes, it is important to consider how the culture conditions might alter DNA integrity and stability and to be able to alter conditions appropriately (Martin et al. 2019).
Growing follicles in vitro
Complete IVG of oocytes from primordial stages with subsequent in vitro maturation of oocytes (IVM) and fertilisation (IVF) has been achieved in the mouse resulting in the production of embryos and live young after embryo transfer (ET) (Eppig & O’Brien 1996, O’Brien et al. 2003). This was achieved using a two-step culture system in which primordial follicles were activated in step one and then growing oocytes developed within oocyte-granulosa cell complexes (OGCs) in step two (Eppig & O’Brien 1996, O’Brien et al. 2003). The first study reported the birth of one mouse derived from IVG oocytes but this animal subsequently developed many abnormalities as an adult although it is not clear if these can be attributed to IVG (Eppig & O’Brien 1996). Further work on the culture system led to improvements in the medium and physical environment resulting in several mouse embryos and offspring from in vitro-grown oocytes (O’Brien et al. 2003). Indeed these reports have been the only ones to date that have demonstrated complete oocyte development using a rodent system from the primordial stage in vitro, indicative of the complexity and technical challenges involved (Eppig & O’Brien 1996, O’Brien et al. 2003). Further studies have recently demonstrated that complete in vitro development of mouse oocytes starting from stages before primordial follicle formation is possible and developmentally competent oocytes capable of being fertilised and forming embryos have been produced from induced pluripotent stem cells (iPSCs) (Hikabe et al. 2016) and primordial germ cells (PGCs) (Morohaku et al. 2016). The studies using rodent models provide proof of principle that it is possible to achieve complete oocyte development in vitro and have highlighted the challenge of translating these systems to support human oocyte development (Herta et al. 2018).
Several culture systems supporting defined stages of human oocyte development have been reported but only a single report describes the growth of human oocytes from primordial stages to maturity within a multi-step culture system (Fig. 2) (McLaughlin et al. 2018). Whilst this system is not yet optimised it highlights the sequence of environments that may mimic the changing niche required to facilitate development of human oocytes in vitro (Anderson & Telfer 2018, McLaughlin et al. 2018).
Steps needed in a multi-step culture system (McLaughlin et al. 2018) to support in vitro growth (IVG) of oocytes starting from human primordial follicles. Step 1: culture of ovarian cortex for 7 days to promote activation of primordial follicles. At the end of the culture period any multi-laminar follicles present are isolated by micro-dissection. Step 2: Isolated follicles are cultured individually in v-shaped wells until antral cavities are formed. Step 3: Oocyte-granulosa cell complexes are removed from intact antral follicles and placed on membranes for up to 4 days. Step 4: Maturation media (IVM) is added to Oocyte-granulosa cell complexes overnight and analysed for the presence of a Metaphase II spindle and a polar body.
Citation: Reproduction 158, 5; 10.1530/REP-19-0077
Supporting initiation/activation of primordial follicles in vitro
Isolated primordial follicles do not activate to grow in vitro. Oktay and colleagues (1997) demonstrated that primordial follicles that were isolated from thawed human cortical tissue were as viable as those isolated from fresh tissue (Oktay et al. 1997) but culturing isolated primordial follicles resulted in poor survival rates (Hovatta et al. 1997, Abir et al. 1999). Therefore culture systems where primordial follicles are the starting stage need to start with pieces of human ovarian cortex rather than isolated primordial follicles (Hovatta et al. 1997, 1999, Picton & Gosden 2000, Telfer et al. 2008, Garor et al. 2009, McLaughlin et al. 2018). For over 20 years there has been the development of systems supporting the initiation of human primordial follicle growth in vitro (Hovatta et al. 1997, 1999, Wright et al. 1999, Picton & Gosden 2000, Hreinsson et al. 2002, Telfer et al. 2008, Garor et al. 2009, Ding et al. 2010, Anderson et al. 2014, McLaughlin et al. 2014, 2018). Different rates of follicle activation have been observed in each of these systems and this appears to be associated with the preparation and architecture of the starting tissue, indicative of the importance of mechanical signalling to cell pathways regulating primordial follicle activation (Shah et al. 2018).
Whilst the activation of primordial follicles in vitro and early follicle development are critical features of an IVG system, the mechanisms regulating follicle activation are still not well understood (Picton & Gosden 2000, Herta et al. 2018). Regulation of activation requires a complex interaction of inhibitory, stimulatory and maintenance factors (Zhang & Liu 2015).
Anti-Mullerian hormone (AMH) produced in the granulosa cells of growing follicles has been shown to have an inhibitory effect on primordial follicle activation in mice (Durlinger et al. 2002) and to attenuate the action of FSH on growing follicles (Durlinger et al. 2001). Human ovarian tissue xenografted to immunodeficient mice with engineered endothelial cells (ECs) to deliver AMH leads to a reduction in primordial follicle activation and growth during a 2-week grafting period (Man et al. 2017). In contrast, human ovarian cortical tissue cultured in the presence of AMH resulted in increased activation and improved development of growing follicles (Schmidt et al. 2005). These results illustrate the differences of action in vitro and in vivo and also of timings and mode of delivery.
Components of the phosphatidylinositol-3’-kinase (PI3K-AKT) signalling pathway within oocytes have been shown to be key regulators of primordial follicle activation in mouse (Reddy et al. 2008, Zhang & Liu 2015), human (Li et al. 2010, McLaughlin et al. 2014, Grosbois & Demeestere 2018) and cow (Maidarti et al. 2019). The PI3K-AKT signalling pathway is the main non-gonadotrophic growth factor signalling pathway regulating growth and differentiation of ovarian follicles (Dupont & Scaramuzzi 2016).
It has been demonstrated that a component of the PI3K pathway, the phosphatase and tensin homolog deleted on chromosome ten (PTEN), negatively regulates this pathway resulting in the suppression of follicle initiation/activation (Reddy et al. 2008). A member of the forkhead transcription factors family (FOXO3a) acts as downstream effector of the PI3K pathway resulting in inhibition of follicle activation (Castrillon et al. 2003). Another component of the PI3K pathway is the mammalian target of rapamycin complex 1 (mTORC1), a regulator of cell growth and proliferation and involved in regulating primordial follicle activation (Adhikari & Liu 2010). Data obtained from knockout mouse models have confirmed that PTEN within the oocyte inhibits follicle activation whilst mTORC1 promotes activation.
Significant activation of primordial follicles occurs when pieces of ovarian cortex are prepared by fragmentation and this appears to be a consequence of disrupting the Hippo signalling pathway (Badouel & McNeill 2011) during tissue preparation (Kawamura et al. 2013, Hsueh et al. 2015, Grosbois & Demeestere 2018). This emphasises the importance of biomechanical forces on these processes (Shah et al. 2018).
Observations on the role of the Hippo signalling pathway may explain differences in results observed between culture systems. Human ovarian cortex prepared as dense cubes in vitro results in low levels of follicle activation (Hovatta et al. 1997, 1999, Wright et al. 1999). In contrast, significant primordial follicle activation occurs in human ovarian cortex prepared as micro-cortex, where underlying stroma is removed and tissue interactions are disrupted (Telfer et al. 2008, McLaughlin et al. 2018). The combination of disruption of the Hippo pathway by fragmentation and manipulation of components of the PI3K pathway (PTEN and mTORC1) results in enhanced activation of follicles (Li et al. 2010, Kawamura et al. 2013, McLaughlin et al. 2014, Hsueh et al. 2015, Grosbois & Demeestere 2018, Maidarti et al. 2019). These observations confirm the importance of tissue preparation in these systems.
The action of PTEN can be inhibited pharmacologically using vanadium compounds such as bisperoxovanadium (Spinelli et al. 2015). Inhibition of PTEN in human tissue prior to transplantation has been used for the treatment of premature ovarian insufficiency (POI) and has led to increased follicle activation (Kawamura et al. 2013). PTEN inhibition in cultured human ovarian cortex also results in more primordial follicles being activated and an increase in the number of secondary follicles that can be isolated (McLaughlin et al. 2014) but further development of these follicles is compromised as demonstrated by poor growth and survival after isolation (McLaughlin et al. 2014, Grosbois & Demeestere 2018). The observed effects on secondary follicles following inhibition of PTEN at the activation stage could be attributed to direct effects on DNA repair response mechanisms within the oocyte/follicle and increased DNA damage (Martin et al. 2019). Primordial follicle activation has been shown to be increased in cultured bovine ovarian cortex where PTEN has been inhibited pharmacologically (Maidarti et al. 2019). This increased activation rate is accompanied by a resultant increase in DNA damage and alterations in DNA repair mechanisms (Maidarti et al. 2019). These deleterious effects are not observed when bovine follicles are activated in media containing activin and FSH without inhibiting PTEN (McLaughlin & Telfer 2010).
Any culture system removes follicles from in vivo control mechanisms (paracrine and endocrine) that will determine their rate of growth, but tissue architecture and stromal density have been shown to have an influence on follicle activation and subsequent growth rate. Follicles within cortical tissue prepared as solid cubes with dense stroma show little growth initiation (Hovatta et al. 1997) when compared with cortex prepared as flattened ‘sheets’, where much of the underlying stroma is removed (Telfer et al. 2008, McLaughlin et al. 2018). How follicles respond to stimulatory and inhibitory factors and therefore their growth rate is influenced by the tissue environment (McLaughlin & McIver 2009). Follicles initiated to grow within cortical tissue can develop to multi-laminar stages, but their growth is inhibited if they are not removed from the cortical tissue (Hovatta et al. 1999, Telfer et al. 2008). Follicles that reach multi-laminar stages start to degenerate if they are not removed from the cortical stromal environment and cultured individually and show a loss of oocyte-somatic cell connections (Telfer et al. 2008, McLaughlin & Telfer 2010, Anderson et al. 2014, McLaughlin et al. 2018).
Growth of multi-laminar follicles in vitro
Isolation of multi-laminar stages from the ovarian cortex can be achieved by using enzymes, mechanical isolation or by combining both methods. Multi-laminar follicles can be isolated from ovarian stroma using a mixture of collagenase and DNase; however, this combination can cause damage resulting in decreased survival of growing follicles (Telfer et al. 2000). The use of liberase for follicle isolation is gentler than collagenase and leads to less damage (Dolmans et al. 2006, Rice et al. 2008). Isolated follicles need to maintain their structure during the second stage of IVG and the presence of theca cells is essential for this. Given that theca cells can be compromised by enzymatic preparations (Wandji et al. 1996, Telfer et al. 2000) it is better to isolate follicles by using needles to remove them from the stroma whilst preserving follicular integrity by maintaining the basal lamina and thecal layers (Telfer et al. 2008, McLaughlin et al. 2018). There are now several culture systems that have been developed to support the growth of isolated multi-laminar (pre-antral) follicles (Telfer & Zelinski 2013).
Tissue engineering has been applied to the challenge of growing multi-laminar structures that could grow up to several millimetres in diameter (Shea et al. 2014, Rajabi et al. 2018a). Human pre-antral follicles can be encapsulated in bio-matrices such as alginate which supports follicle structure and growth in vitro (Xu et al. 2009). Alginate is a polysaccharide that is found in the cell wall of algae and forms a gel which follicles can be embedded in. Alginate forms controlled set gels with ridgidity that can be manipulated. It is important to balance flexibility of the gel to accommodate cell proliferation with its rigidity to avoid collapse of the follicle structure (Skory et al. 2015). The degree of rigidity affects follicle development as demonstrated by reports that show poor growth of mouse follicles embedded in 1% alginate gels (Heise et al. 2005) whilst 0.5% gels support the development of fully grown human oocytes (Xu et al. 2009). These studies demonstrate that the physical environment of the ovary is a regulating force and that biomechanical signals can be mimicked by an appropriate support matrix (West-Farrell et al. 2009).
Other scaffolds such as de-cellularised ovarian tissue (Laronda et al. 2015) and 3D microporous scaffolds are being developed as matrices and are being applied to support human pre-antral follicle growth (Laronda et al. 2017, Pors et al. 2019). Electrospun patterned porous scaffolds are being explored to support follicle development and these may be more accessible and reproducible than de-cellularised tissue (Liverani et al. 2019).
No matrices or scaffolds are utilised in the multi-step culture system developed for human follicles (McLaughlin et al. 2018). During step 2 of the multi-step system isolated follicles are placed free floating in v-shaped micro-well plates. This system supports follicular structure in vitro as well as growth, differentiation and antral formation during a relatively short culture period of up to 10 days (Telfer et al. 2008, McLaughlin et al. 2018).
Following initiation of follicle growth within the strip, follicles develop to multi-laminar stages but do not survive within the cortical environment (Hovatta et al. 1997, Telfer et al. 2008). Therefore follicles need to be released from the cortical stromal environment and cultured individually (Telfer et al. 2008, McLaughlin et al. 2018). Pre-antral follicles isolated from ovarian cortex and grown in vitro undergo significant growth and development and whilst this can be rapid it represents growth without brakes since the follicle has been removed from all regulatory mechanisms. Two approaches have been taken by researchers developing culture systems to support the growth of pre-antral follicle, one is a ‘slow’ grow system where isolated follicles are grown for protracted periods in an attempt to mimic growth rates in vivo (Chambers et al. 2010). Multi-laminar follicles isolated enzymatically from fresh human ovarian tissue can undergo complete oocyte growth within 30 days (Xu et al. 2009) and some are capable of meiotic maturation (Xiao et al. 2015). Whilst these systems may be presented as slow growth systems they still support a more rapid development than that observed in vivo.
Developing pre-antral human follicles to pre-ovulatory diameters present technical challenges but oocyte development may not require the expansion of follicle size associated with pre-ovulatory stages and so a ‘fast’ growth system that focuses on oocyte development has been developed. Follicles activated to grow within fragments of ovarian cortex reach multi-laminar stages within 7 days and when they are removed from the cortex and cultured individually achieve antral formation and significant oocyte growth within a further 7–10 days of in vitro growth in the presence of activin A and FSH (Telfer et al. 2008, McLaughlin et al. 2018).
Developmentally competent mouse oocytes can be obtained from primordial stages using a two-step culture system where follicles are activated within ovarian fragments and then oocyte-granulosa cell complexes (OGCs) are isolated and cultured during step 2 (O’Brien et al. 2003). This system does not require antral follicles to be formed but by maintaining oocyte-somatic cell connections within oocyte–granulosa cell complexes (OGCs) complete oocyte development can be supported (O’Brien et al. 2003). The mouse system has informed the development of step three of the human multi-step culture system (McLaughlin et al. 2018). Whilst isolated human follicles cultured individually form antral cavities within 6–8 days, rather than attempt to maintain the development of large antral follicles, oocyte–granulosa cell complexes are removed from the follicle for further growth and development. The removal of complexes at this stage is technically demanding and is achieved by applying gentle pressure to the antral follicle (Telfer et al. 2008, McLaughlin et al. 2018). Intact complexes with complete cumulus and adherent mural granulosa cells are selected for further growth in step 3 of the multi-step system (Fig. 2). These complexes can produce oocytes of at least 100 microns in the presence of activin-A and rhFSH, and some of these can reach metaphase II (McLaughlin et al. 2018).
Optimising conditions to support activation and early growth
A multitude of factors affect follicle/oocyte development in vivo, either by promoting growth or degeneration. There is still a lack of knowledge on the exact requirements to support in vitro growth therefore optimising conditions has been extremely difficult (Telfer & Zelinski 2013, Rajabi et al. 2018b). Early culture systems to support activation and growth used medium with serum but defined media containing human serum albumin (HSA) and a combination of insulin, selenium and transferrin (ITS) is now being used (Telfer et al. 2008, McLaughlin et al. 2018). Several types of basal culture media including MEM alpha, Waymouths medium and McCoys 5A have been used with our preference being for McCoys 5A (Telfer et al. 2008, McLaughlin et al. 2018). An essential component of the culture media is ascorbic acid which is added fresh at each media change to reduce cell death (Thomas et al. 2001). The activation and growth of primordial follicles occurs in the presence of basic medium, and this process can be inhibited by the addition of growth factors such as activin (Ding et al. 2010). A greater understanding of how specific factors influence the process of activation is essential and will lead to improvements in number and quality of IVG follicles. Another approach that is being developed to increase the number of follicles activated in vitro is the manipulation of key signalling pathways controlling this process. As discussed above the PI3K/PTEN/Akt pathway has a major role in activation of primordial follicles but balancing increasing activation by manipulating this pathway with possible damage and effects on DNA repair mechanisms needs to be determined (Maidarti et al. 2019).
The addition of several growth factors and hormones has been tested in a range of species but there are conflicting reports and still no consensus on the timing and concentration of key additives (Rajabi et al. 2018b). It is well recognised that oocyte–somatic cell interactions need to be maintained to support oocyte development. Cell–cell communication is affected by members of the TGF beta superfamily, in particular oocyte-specific factors, growth differentiation factor-9 (GDF-9) and bone morphogenetic protein 15 (BMP-15) (Sanfins et al. 2018). Human GDF-9 and BMP 15 act in a synergistic fashion to promote cell proliferation and cell–cell interaction (Reader et al. 2016). In humans GDF-9 and BMP-15 is expressed in the oocyte and cumulus granulosa cells and increased mRNA levels in cumulus cells has been correlated with an increase in oocyte maturation and subsequent fertilisation rates in human (Li et al. 2014). The addition of either GDF-9 or BMP-15 to cultured human ovarian cortex can increase the activation of primordial follicles with BMP-15 having more beneficial effects on early growth (Kedem et al. 2011).
Successful growth of multi-laminar follicles in step 2 of the multi-step system is dependent on maintaining oocyte–somatic cell interactions during this stage (Li & Albertini 2013). Experiments using human tissue demonstrated that the addition of activin resulted in antral cavity formation in vitro (Telfer et al. 2008) and the combination of activin and a low-dose FSH during stage 2 of growth maintains oocyte–somatic cell interactions leading to improved health of multi-laminar stages and antral formation in a bovine model (McLaughlin et al. 2010) and has been successfully applied to a human system (McLaughlin et al. 2018). Given the range and complexity of factors involved in oocyte/follicle development it is clear that the concentration, mode of delivery and timings of exposure to specific factors need to be determined. Co-ordinating such a complex system will require the development of systems that can facilitate a dynamic culture system. Researchers are working on bioreactor systems to regulate timings, concentration and oxygen availability in a dynamic process (Catapano et al. 2019).
Since oocyte size indicates its ability to resume meiosis (Fair et al. 1995, Sirard 2011) complete in vitro development systems must support oocyte growth rather than follicle growth (McLaughlin et al. 2018). Differences in growth rate of oocytes have been observed between those cultured within intact follicles for the entire culture period (Xu et al. 2009, Xiao et al. 2015) and those that are removed as complexes after an antral cavity has formed and cultured on membranes (Telfer et al. 2008, McLaughlin et al. 2018). Comparisons of culture systems with different growth rates is required to determine optimal conditions. It will be essential to determine whether in vitro growth is capable of supporting the development of healthy human oocytes. Any deleterious effects on oocyte function and epigenetic changes are as yet unknown and will require a large animal model such as cow or sheep to determine the safety of these techniques. In addition, metabolic studies such as those carried out on sheep follicles over a prolonged culture period should be conducted for human follicles to determine how growth period affects metabolism as an indicator of oocyte health and competence (Harris et al. 2010, Hemmings et al. 2012).
Meiotic maturation of oocytes from IVG human follicles
Once IVG oocytes are fully grown they need to be placed in maturation media (IVM) to determine their capacity to resume meiosis and reach metaphase II (Fig. 2). IVM has been successfully utilised in domestic species such as cows (Van den Hurk et al. 2000, Hirao et al. 2013, Dieci et al. 2016) and pigs (Hirao et al. 1994) for embryo production. IVM of human oocytes has been applied but success rates have been variable (Nogueira et al. 2012, Chian et al. 2013). Whilst IVM of immature human oocytes resulted in a live birth in 1991 (Cha et al. 1991), this technique was being used whilst IVF was being developed (Edwards et al. 1969). IVM is now used clinically in a small number of centres with variable success rates (Chian et al. 2013). Success rates are related to the source of oocytes selected for IVM with it being well recognised that successful maturation of immature oocytes is lower than that of oocytes obtained from stimulated ovaries (Nogueira et al. 2012, Chian et al. 2013). Many of the oocytes harvested for IVM will be intrinsically unable to undergo maturation therefore optimisation of IVM conditions is also required if IVG is ever to be applied clinically (Shirasawa & Terada 2017).
Some oocytes that survive the multi-step culture system are capable of meiotic maturation following an IVM protocol (McLaughlin et al. 2018). Whilst some IVG oocytes form metaphase II spindles their polar bodies are larger than normal (McLaughlin et al. 2018). It is not known what causes the formation of large bodies in IVG-derived oocytes, but polar body size can be affected by the position of the spindle in relation to the oocyte cortex (Barrett & Albertini 2010). It has been shown that large polar bodies are extruded from human oocytes undergoing IVM when spindle contact with the oocyte cortex is lost (Coticchio et al. 2013). The goal of any IVG system is to support the development of oocytes that are competent to be fertilised and are epigenetically normal. Research focusing on optimising each of the stages of IVG is required, and it is essential to determine the epigenetic status of IVG oocytes and of any embryos that might be formed (Anckaert et al. 2013).
Steps towards clinical translation
The main clinical application of IVG is within fertility preservation therefore it is important to determine whether cryopreserved/thawed ovarian tissue can be cultured. Human follicles isolated from cryopreserved tissue have been shown to be viable and the birth rate from tissue re-implantation is now more than 130 babies (Donnez & Dolmans 2017). Analysis of frozen thawed tissue indicates high viability of follicles (Kristensen et al. 2018) and activation in vitro of primordial follicles in frozen thawed ovarian cortex can be comparable to fresh tissue (Grosbois & Demeestere 2018). Whilst follicles isolated from frozen thawed tissue after activation and culture appear to grow in a similar way detailed testing is still required.
Future developments: making new oocytes from stem cells
The starting material for the in vitro growth systems described above is the primordial follicle that has been formed in vivo before birth. There is now a focus on trying to produce in vitro-derived oocytes, from sources such as embryonic stem cells (ESCs), induced pluripotent stem cells (iPSCs) or putative oogonial stem cells (OSCs). The ability to produce oocytes in this way would provide women that have few or no oocytes with the possibility of genetically related offspring and would alleviate the need for donor eggs (Ilic et al. 2019). Several studies have shown that putative OSCs can be isolated from ovarian cortex of adult women (White et al. 2012, Anderson & Telfer 2018, Clarkson et al. 2018, Silvestris et al. 2018). These cells have the capacity to form what appear to be oocytes when they are grown in vitro and provided with a somatic cell support (White et al. 2012, Clarkson et al. 2018, Silvestris et al. 2018). Controversy has been generated over the biological significance of these cells and this remains unknown but the identification and isolation of populations of cells from the adult human ovary that have germ line potential is a significant finding. These cells could have the potential to improve infertility treatments if new oocytes can be formed and grown in vitro (Anderson & Telfer 2018). Whilst the production of oocytes from stem cells is promising, it is clear that some cells under certain conditions can form morphological oocytes, but these do not enter meiosis. The process of oocyte differentiation can be dissociated from meiosis, so if functional artificial oocytes are to be obtained, then it will be essential to understand the connection between oocyte differentiation and entry into meiosis (Handel et al. 2014).
The development of functional oocytes from stem cells is dependent on the methodologies discussed in this review to grow human immature oocytes from primordial stages to mature oocytes entirely in vitro. As highlighted here there are many practical and conceptual obstacles remaining before clinical application of any of these technologies can ever be fully realised.
Summary
The practice of fertility preservation is a new and fast developing field. Cryopreservation and re-implantation of ovarian tissue developed by Roger Gosden laid the foundations and IVG systems would be an important addition. The capacity to successfully develop human oocytes from immature stages through to maturation and fertilisation in vitro would offer an alternative to autologous transplantation. In addition it has the potential to be the next generation IVF. Proof of concept that complete in vitro growth of human oocytes is possible has been achieved but optimisation is now required (McLaughlin et al. 2018). There is an overarching need to avail models of human gametogenesis under conditions that will allow for detailed analyses of the sequential stages of oogenesis in the human. By culturing human oocytes/follicles we have gained greater insight to the process of human oogenesis and better understanding of human oocyte development.
There are many women for whom current methods of fertility preservation are unsuitable and the process of IVG would be beneficial to them but cautious optimism needs to be tempered with realism. There is a long road ahead before optimisation and thorough testing can be carried out to demonstrate the safety of these techniques (Anderson & Telfer 2018).
Declaration of interest
The author declares that there is no conflict of interest that could be perceived as prejudicing the impartiality of the research reported.
Funding
Work in this area carried out in Evelyn Telfer’s laboratory has been supported by MRC grants GO901838, MR/L00299X/1, MR/R003246/1.
Acknowledgements
E E T acknowledges the assistance of Dr Yvonne Clarkson during the preparation of this manuscript.
References
Abir R, Roizman P, Fisch B, Nitke S, Okon E, Orvieto R & Ben Rafael Z 1999 Pilot study of isolated early human follicles cultured in collagen gels for 24 hours. Human Reproduction 1299–1301. (https://doi.org/10.1093/humrep/14.5.1299)
Adhikari D & Liu K 2010 MTOR signaling in the control of activation of primordial follicles. Cell Cycle 1673–1674. (https://doi.org/10.4161/cc.9.9.11626)
Anckaert E, De Rycke M & Smitz J 2013 Culture of oocytes and risk of imprinting defects. Human Reproduction Update 52–66. (https://doi.org/10.1093/humupd/dms042)
Anderson RA & Baird DT 2019 The development of ovarian tissue cryopreservation in Edinburgh: translation from a rodent model through validation in a large mammal and then into clinical practice. Acta Obstetricia et Gynecologica Scandinavica 545–549. (https://doi.org/10.1111/aogs.13560)
Anderson RA, McLaughlin M, Wallace WH, Albertini DF & Telfer EE 2014 The immature human ovary shows loss of abnormal follicles and increasing follicle developmental competence through childhood and adolescence. Human Reproduction 97–106. (https://doi.org/10.1093/humrep/det388)
Anderson RA, Mitchell RT, Kelsey TW, Spears N, Telfer EE & Wallace WH 2015 Cancer treatment and gonadal function: experimental and established strategies for fertility preservation in children and young adults. Lancet. Diabetes & Endocrinology 556–567. (https://doi.org/10.1016/S2213-8587(15)00039-X)
Anderson RA & Telfer EE 2018 Being a good egg in the 21st century. British Medical Bulletin 83–89. (https://doi.org/10.1093/bmb/ldy023)
Anderson RA, Wallace WHB & Telfer EE 2017 Ovarian tissue cryopreservation for fertility preservation: clinical and research perspectives. Human Reproduction Open hox001 . (https://doi.org/10.1093/hropen/hox001)
Badouel C & McNeill H 2011 SnapShot: the hippo signaling pathway. Cell 484–484.e1. (https://doi.org/10.1016/j.cell.2011.04.009)
Barrett SL & Albertini DF 2010 Cumulus cell contact during oocyte maturation in mice regulates meiotic spindle positioning and enhances developmental competence. Journal of Assisted Reproduction and Genetics 29–39. (https://doi.org/10.1007/s10815-009-9376-9)
Castrillon DH, Miao L, Kollipara R, Horner JW & DePinho RA 2003 Suppression of ovarian follicle activation in mice by the transcription factor Foxo3a. Science 215–218. (https://doi.org/10.1126/science.1086336)
Catapano G, Fragomeni G, Falvo D’Urso Labate G, De Napoli L, Barbato V, Di Nardo M, Costanzo V, Capriglione T, Gualtieri R & Talevi R 2019 Do bioreactor designs with more efficient oxygen supply to ovarian cortical tissue fragments enhance follicle viability and growth in vitro? Processes 450 . (https://doi.org/10.3390/pr7070450)
Cha KY, Koo JJ, Ko JJ, Choi DH, Han SY & Yoon TK 1991 Pregnancy after in vitro fertilization of human follicular oocytes collected from nonstimulated cycles, their culture in vitro and their transfer in a donor oocyte program. Fertility and Sterility 109–113. (https://doi.org/10.1016/s0015-0282(16)54068-0)
Chambers EL, Gosden RG, Yap C & Picton HM 2010 In situ identification of follicles in ovarian cortex as a tool for quantifying follicle density, viability and developmental potential in strategies to preserve female fertility. Human Reproduction 2559–2568. (https://doi.org/10.1093/humrep/deq192)
Chian RC, Uzelac PS & Nargund G 2013 In vitro maturation of human immature oocytes for fertility preservation. Fertility and Sterility 1173–1181. (https://doi.org/10.1016/j.fertnstert.2013.01.141)
Clarkson YL, McLaughlin M, Waterfall M, Dunlop CE, Skehel PA, Anderson RA & Telfer EE 2018 Initial characterisation of adult human ovarian cell populations isolated by DDX4 expression and aldehyde dehydrogenase activity. Scientific Reports 6953 . (https://doi.org/10.1038/s41598-018-25116-1)
Coticchio G, Guglielmo MC, Dal Canto M, Fadini R, Mignini Renzini M, De Ponti E, Brambillasca F & Albertini DF 2013 Mechanistic foundations of the metaphase II spindle of human oocytes matured in vivo and in vitro. Human Reproduction 3271–3282. (https://doi.org/10.1093/humrep/det381)
Dieci C, Lodde V, Labreque R, Dufort I, Tessaro I, Sirard MA & Luciano AM 2016 Differences in cumulus cell gene expression indicate the benefit of a pre-maturation step to improve in-vitro bovine embryo production. Molecular Human Reproduction 882–897. (https://doi.org/10.1093/molehr/gaw055)
Ding CC, Thong KJ, Krishna A & Telfer EE 2010 Activin A inhibits activation of human primordial follicles in vitro. Journal of Assisted Reproduction and Genetics 141–147. (https://doi.org/10.1007/s10815-010-9395-6)
Dolmans MM & Amorim CA 2019 Construction and use of artificial ovaries. Reproduction 158 F15–F25. (https://doi.org/10.1530/REP-18-0536)
Dolmans MM, Michaux N, Camboni A, Martinez-Madrid B, Van Langendonckt A, Nottola SA & Donnez J 2006 Evaluation of Liberase, a purified enzyme blend, for the isolation of human primordial and primary ovarian follicles. Human Reproduction 413–420. (https://doi.org/10.1093/humrep/dei320)
Donnez J & Dolmans MM 2017 Fertility preservation in women. The New England Journal of Medicine 1657–1665. (https://doi.org/10.1056/NEJMra1614676)
Dupont J & Scaramuzzi RJ 2016 Insulin signalling and glucose transport in the ovary and ovarian function during the ovarian cycle. The Biochemical Journal 1483–1501. (https://doi.org/10.1042/BCJ20160124)
Durlinger AL, Gruijters MJ, Kramer P, Karels B, Kumar TR, Matzuk MM, Rose UM, de Jong FH, Uilenbroek JT & Grootegoed JA, et al.2001 Anti-Mullerian hormone attenuates the effects of FSH on follicle development in the mouse ovary. Endocrinology 4891–4899. (https://doi.org/10.1210/endo.142.11.8486)
Durlinger AL, Visser JA & Themmen AP 2002 Regulation of ovarian function: the role of anti-Mullerian hormone. Reproduction 601–609. (https://doi.org/10.1530/rep.0.1240601)
Edwards RG, Bavister BD & Steptoe PC 1969 Early stages of fertilization in vitro of human oocytes matured in vitro. Nature 632–635. (https://doi.org/10.1038/221632a0)
Eppig JJ & O’Brien MJ 1996 Development in vitro of mouse oocytes from primordial follicles. Biology of Reproduction 197–207. (https://doi.org/10.1095/biolreprod54.1.197)
Fair T, Hyttel P & Greve T 1995 Bovine oocyte diameter in relation to maturational competence and transcriptional activity. Molecular Reproduction and Development 437–442. (https://doi.org/10.1002/mrd.1080420410)
Garor R, Abir R, Erman A, Felz C, Nitke S & Fisch B 2009 Effects of basic fibroblast growth factor on in vitro development of human ovarian primordial follicles. Fertility and Sterility 1967–1975. (https://doi.org/10.1016/j.fertnstert.2008.04.075)
Grosbois J & Demeestere I 2018 Dynamics of PI3K and Hippo signaling pathways during in vitro human follicle activation. Human Reproduction 1705–1714. (https://doi.org/10.1093/humrep/dey250)
Handel MA, Eppig JJ & Schimenti JC 2014 Applying “gold standards” to in-vitro-derived germ cells. Cell 1257–1261. (https://doi.org/10.1016/j.cell.2014.05.019)
Harris SE, Maruthini D, Tang T, Balen AH & Picton HM 2010 Metabolism and karyotype analysis of oocytes from patients with polycystic ovary syndrome. Human Reproduction 2305–2315. (https://doi.org/10.1093/humrep/deq181)
Heise M, Koepsel R, Russell AJ & McGee EA 2005 Calcium alginate microencapsulation of ovarian follicles impacts FSH delivery and follicle morphology. Reproductive Biology and Endocrinology : RB&E 47 . (https://doi.org/10.1186/1477-7827-3-47)
Hemmings KE, Leese HJ & Picton HM 2012 Amino acid turnover by bovine oocytes provides an index of oocyte developmental competence in vitro. Biology of Reproduction 165, 1–165,12. (https://doi.org/10.1095/biolreprod.111.092585)
Herta AC, Lolicato F & Smitz JEJ 2018 In vitro follicle culture in the context of IVF. Reproduction 156 F59–F73. (https://doi.org/10.1530/REP-18-0173)
Hikabe O, Hamazaki N, Nagamatsu G, Obata Y, Hirao Y, Hamada N, Shimamoto S, Imamura T, Nakashima K & Saitou M, et al.2016 Reconstitution in vitro of the entire cycle of the mouse female germ line. Nature 299–303. (https://doi.org/10.1038/nature20104)
Hirao Y, Nagai T, Kubo M, Miyano T, Miyake M & Kato S 1994 In vitro growth and maturation of pig oocytes. Journal of Reproduction and Fertility 333–339. (https://doi.org/10.1530/jrf.0.1000333)
Hirao Y, Naruse K, Kaneda M, Somfai T, Iga K, Shimizu M, Akagi S, Cao F, Kono T & Nagai T, et al.2013 Production of fertile offspring from oocytes grown in vitro by nuclear transfer in cattle. Biology of Reproduction 57 . (https://doi.org/10.1095/biolreprod.113.109439)
Hovatta O, Silye R, Abir R, Krausz T & Winston RM 1997 Extracellular matrix improves survival of both stored and fresh human primordial and primary ovarian follicles in long-term culture. Human Reproduction 1032–1036. (https://doi.org/10.1093/humrep/12.5.1032)
Hovatta O, Wright C, Krausz T, Hardy K & Winston RM 1999 Human primordial, primary and secondary ovarian follicles in long-term culture: effect of partial isolation. Human Reproduction 2519–2524. (https://doi.org/10.1093/humrep/14.10.2519)
Hreinsson JG, Scott JE, Rasmussen C, Swahn ML, Hsueh AJ & Hovatta O 2002 Growth differentiation factor-9 promotes the growth, development, and survival of human ovarian follicles in organ culture. The Journal of Clinical Endocrinology and Metabolism 316–321. (https://doi.org/10.1210/jcem.87.1.8185)
Hsueh AJ, Kawamura K, Cheng Y & Fauser BC 2015 Intraovarian control of early folliculogenesis. Endocrine Reviews 1–24. (https://doi.org/10.1210/er.2015.36.issue-1.edboard)
Ilic D, Telfer EE, Ogilvie C, Kolundzic N & Khalaf Y 2019 What can stem cell technology offer to IVF patients? BJOG : an International Journal of Obstetrics and Gynaecology 824–827. (https://doi.org/10.1111/1471-0528.15638)
Kawamura K, Cheng Y, Suzuki N, Deguchi M, Sato Y, Takae S, Ho CH, Kawamura N, Tamura M & Hashimoto S, et al.2013 Hippo signaling disruption and Akt stimulation of ovarian follicles for infertility treatment. Proceedings of the National Academy of Sciences of the United States of America 17474–17479. (https://doi.org/10.1073/pnas.1312830110)
Kedem A, Fisch B, Garor R, Ben-Zaken A, Gizunterman T, Felz C, Ben-Haroush A, Kravarusic D & Abir R 2011 Growth differentiating factor 9 (GDF9) and bone morphogenetic protein 15 both activate development of human primordial follicles in vitro, with seemingly more beneficial effects of GDF9. The Journal of Clinical Endocrinology and Metabolism 96 E1246–E1254. (https://doi.org/10.1210/jc.2011-0410)
Kristensen SG, Liu Q, Mamsen LS, Greve T, Pors SE, Bjorn AB, Ernst E, Macklon KT & Andersen CY 2018 A simple method to quantify follicle survival in cryopreserved human ovarian tissue. Human Reproduction 2276–2284. (https://doi.org/10.1093/humrep/dey318)
Laronda MM, Jakus AE, Whelan KA, Wertheim JA, Shah RN & Woodruff TK 2015 Initiation of puberty in mice following decellularized ovary transplant. Biomaterials 20–29. (https://doi.org/10.1016/j.biomaterials.2015.01.051)
Laronda MM, Rutz AL, Xiao S, Whelan KA, Duncan FE, Roth EW, Woodruff TK & Shah RN 2017 A bioprosthetic ovary created using 3D printed microporous scaffolds restores ovarian function in sterilized mice. Nature Communications 15261 . (https://doi.org/10.1038/ncomms15261)
Li J, Kawamura K, Cheng Y, Liu S, Klein C, Liu S, Duan EK & Hsueh AJ 2010 Activation of dormant ovarian follicles to generate mature eggs. Proceedings of the National Academy of Sciences of the United States of America 10280–10284. (https://doi.org/10.1073/pnas.1001198107)
Li R & Albertini DF 2013 The road to maturation: somatic cell interaction and self-organization of the mammalian oocyte. Nature Reviews. Molecular Cell Biology 141–152. (https://doi.org/10.1038/nrm3531)
Li Y, Li RQ, Ou SB, Zhang NF, Ren L, Wei LN, Zhang QX & Yang DZ 2014 Increased GDF9 and BMP15 mRNA levels in cumulus granulosa cells correlate with oocyte maturation, fertilization, and embryo quality in humans. Reproductive Biology and Endocrinology: RB&E 81 . (https://doi.org/10.1186/1477-7827-12-81)
Liverani L, Raffel N, Fattahi A, Preis A, Hoffmann I, Boccaccini AR, Beckmann MW & Dittrich R 2019 Electrospun patterned porous scaffolds for the support of ovarian follicles growth: a feasibility study. Scientific Reports 1150 . (https://doi.org/10.1038/s41598-018-37640-1)
Maidarti M, Clarkson YL, McLaughlin M, Anderson RA & Telfer EE 2019 Inhibition of PTEN activates bovine non-growing follicles in vitro but increases DNA damage and reduces DNA repair response. Human Reproduction 297–307. (https://doi.org/10.1093/humrep/dey354)
Man L, Park L, Bodine R, Ginsberg M, Zaninovic N, Man OA, Schattman G, Rosenwaks Z & James D 2017 Engineered endothelium provides angiogenic and paracrine stimulus to grafted human ovarian tissue. Scientific Reports 8203 . (https://doi.org/10.1038/s41598-017-08491-z)
Martin JH, Aitken RJ, Bromfield EG & Nixon B 2019 DNA damage and repair in the female germline: contributions to ART. Human Reproduction Update 180–201. (https://doi.org/10.1093/humupd/dmy040)
McLaughlin EA & McIver SC 2009 Awakening the oocyte: controlling primordial follicle development. Reproduction 1–11. (https://doi.org/10.1530/REP-08-0118)
McLaughlin M, Albertini DF, Wallace WHB, Anderson RA & Telfer EE 2018 Metaphase II oocytes from human unilaminar follicles grown in a multi-step culture system. Molecular Human Reproduction 135–142. (https://doi.org/10.1093/molehr/gay002)
McLaughlin M, Bromfield JJ, Albertini DF & Telfer EE 2010 Activin promotes follicular integrity and oogenesis in cultured pre-antral bovine follicles. Molecular Human Reproduction 644–653. (https://doi.org/10.1093/molehr/gaq021)
McLaughlin M, Kinnell HL, Anderson RA & Telfer EE 2014 Inhibition of phosphatase and tensin homologue (PTEN) in human ovary in vitro results in increased activation of primordial follicles but compromises development of growing follicles. Molecular Human Reproduction 736–744. (https://doi.org/10.1093/molehr/gau037)
McLaughlin M & Telfer EE 2010 Oocyte development in bovine primordial follicles is promoted by activin and FSH within a two-step serum-free culture system. Reproduction 971–978. (https://doi.org/10.1530/REP-10-0025)
Morohaku K, Tanimoto R, Sasaki K, Kawahara-Miki R, Kono T, Hayashi K, Hirao Y & Obata Y 2016 Complete in vitro generation of fertile oocytes from mouse primordial germ cells. Proceedings of the National Academy of Sciences of the United States of America 9021–9026. (https://doi.org/10.1073/pnas.1603817113)
Niederberger C, Pellicer A, Cohen J, Gardner DK, Palermo GD, O’Neill CL, Chow S, Rosenwaks Z, Cobo A & Swain JE, et al.2018 Forty years of IVF. Fertility and Sterility 185–324.e5. (https://doi.org/10.1016/j.fertnstert.2018.06.005)
Nogueira D, Sadeu JC & Montagut J 2012 In vitro oocyte maturation: current status. Seminars in Reproductive Medicine 199–213. (https://doi.org/10.1055/s-0032-1311522)
O’Brien MJ, Pendola JK & Eppig JJ 2003 A revised protocol for in vitro development of mouse oocytes from primordial follicles dramatically improves their developmental competence. Biology of Reproduction 1682–1686. (https://doi.org/10.1095/biolreprod.102.013029)
Oktay K, Nugent D, Newton H, Salha O, Chatterjee P & Gosden RG 1997 Isolation and characterization of primordial follicles from fresh and cryopreserved human ovarian tissue. Fertility and Sterility 481–486. (https://doi.org/10.1016/s0015-0282(97)80073-8)
Picton HM & Gosden RG 2000 In vitro growth of human primordial follicles from frozen-banked ovarian tissue. Molecular and Cellular Endocrinology 27–35. (https://doi.org/10.1016/s0303-7207(00)00294-x)
Picton HM, Harris SE, Muruvi W & Chambers EL 2008 The in vitro growth and maturation of follicles. Reproduction 703–715. (https://doi.org/10.1530/REP-08-0290)
Pors SE, Ramlose M, Nikiforov D, Lundsgaard K, Cheng J, Andersen CY & Kristensen SG 2019 Initial steps in reconstruction of the human ovary: survival of pre-antral stage follicles in a decellularized human ovarian scaffold. Human Reproduction 34 1523–1535. (https://doi.org/10.1093/humrep/dez077)
Rajabi Z, Aliakbari F & Yazdekhasti H 2018a Female fertility preservation, clinical and experimental options. Journal of Reproduction & Infertility 125–132.
Rajabi Z, Khokhar Z & Yazdekhasti H 2018b The growth of preantral follicles and the impact of different supplementations and circumstances: a review study with focus on bovine and human preantral follicles. Cellular Reprogramming 164–177. (https://doi.org/10.1089/cell.2017.0068)
Reader KL, Mottershead DG, Martin GA, Gilchrist RB, Heath DA, McNatty KP & Juengel JL 2016 Signalling pathways involved in the synergistic effects of human growth differentiation factor 9 and bone morphogenetic protein 15. Reproduction, Fertility, and Development 491–498. (https://doi.org/10.1071/RD14099)
Reddy P, Liu L, Adhikari D, Jagarlamudi K, Rajareddy S, Shen Y, Du C, Tang W, Hamalainen T & Peng SL, et al.2008 Oocyte-specific deletion of Pten causes premature activation of the primordial follicle pool. Science 611–613. (https://doi.org/10.1126/science.1152257)
Rice S, Ojha K & Mason H 2008 Human ovarian biopsies as a viable source of pre-antral follicles. Human Reproduction 600–605. (https://doi.org/10.1093/humrep/dem390)
Sanfins A, Rodrigues P & Albertini DF 2018 GDF-9 and BMP-15 direct the follicle symphony. Journal of Assisted Reproduction and Genetics 1741–1750. (https://doi.org/10.1007/s10815-018-1268-4)
Schmidt KL, Kryger-Baggesen N, Byskov AG & Andersen CY 2005 Anti-Mullerian hormone initiates growth of human primordial follicles in vitro. Molecular and Cellular Endocrinology 87–93. (https://doi.org/10.1016/j.mce.2004.12.010)
Shah JS, Sabouni R, Cayton Vaught KC, Owen CM, Albertini DF & Segars JH 2018 Biomechanics and mechanical signaling in the ovary: a systematic review. Journal of Assisted Reproduction and Genetics 1135–1148. (https://doi.org/10.1007/s10815-018-1180-y)
Shea LD, Woodruff TK & Shikanov A 2014 Bioengineering the ovarian follicle microenvironment. Annual Review of Biomedical Engineering 29–52. (https://doi.org/10.1146/annurev-bioeng-071813-105131)
Shirasawa H & Terada Y 2017 In vitro maturation of human immature oocytes for fertility preservation and research material. Reproductive Medicine and Biology 258–267. (https://doi.org/10.1002/rmb2.12042)
Silvestris E, Cafforio P, D’Oronzo S, Felici C, Silvestris F & Loverro G 2018 In vitro differentiation of human oocyte-like cells from oogonial stem cells: single-cell isolation and molecular characterization. Human Reproduction 464–473. (https://doi.org/10.1093/humrep/dex377)
Sirard MA 2011 Follicle environment and quality of in vitro matured oocytes. Journal of Assisted Reproduction and Genetics 483–488. (https://doi.org/10.1007/s10815-011-9554-4)
Skory RM, Xu Y, Shea LD & Woodruff TK 2015 Engineering the ovarian cycle using in vitro follicle culture. Human Reproduction 1386–1395. (https://doi.org/10.1093/humrep/dev052)
Spinelli L, Lindsay YE & Leslie NR 2015 PTEN inhibitors: an evaluation of current compounds. Advances in Biological Regulation 102–111. (https://doi.org/10.1016/j.jbior.2014.09.012)
Telfer EE 2019 Future developments: in vitro growth (IVG) of human ovarian follicles. Acta Obstetricia et Gynecologica Scandinavica 653–658. (https://doi.org/10.1111/aogs.13592)
Telfer EE, Binnie JP, McCaffery FH & Campbell BK 2000 In vitro development of oocytes from porcine and bovine primary follicles. Molecular and Cellular Endocrinology 117–123. (https://doi.org/10.1016/s0303-7207(00)00216-1)
Telfer EE & McLaughlin M 2011 In vitro development of ovarian follicles. Seminars in Reproductive Medicine 15–23. (https://doi.org/10.1055/s-0030-1268700)
Telfer EE, McLaughlin M, Ding C & Thong KJ 2008 A two-step serum-free culture system supports development of human oocytes from primordial follicles in the presence of activin. Human Reproduction 1151–1158. (https://doi.org/10.1093/humrep/den070)
Telfer EE & Zelinski MB 2013 Ovarian follicle culture: advances and challenges for human and nonhuman primates. Fertility and Sterility 1523–1533. (https://doi.org/10.1016/j.fertnstert.2013.03.043)
Thomas FH, Leask R, Srsen V, Riley SC, Spears N & Telfer EE 2001 Effect of ascorbic acid on health and morphology of bovine preantral follicles during long-term culture. Reproduction 487–495. (https://doi.org/10.1530/rep.0.1220487)
Van den Hurk R, Abir R, Telfer EE & Bevers MM 2000 Primate and bovine immature oocytes and follicles as sources of fertilizable oocytes. Human Reproduction Update 457–474. (https://doi.org/10.1093/humupd/6.5.457)
Wandji SA, Eppig JJ & Fortune JE 1996 FSH and growth factors affect the growth and endocrine function in vitro of granulosa cells of bovine preantral follicles. Theriogenology 817–832. (https://doi.org/10.1016/0093-691x(96)00011-8)
West-Farrell ER, Xu M, Gomberg MA, Chow YH, Woodruff TK & Shea LD 2009 The mouse follicle microenvironment regulates antrum formation and steroid production: alterations in gene expression profiles. Biology of Reproduction 432–439. (https://doi.org/10.1095/biolreprod.108.071142)
White YA, Woods DC, Takai Y, Ishihara O, Seki H & Tilly JL 2012 Oocyte formation by mitotically active germ cells purified from ovaries of reproductive-age women. Nature Medicine 413–421. (https://doi.org/10.1038/nm.2669)
Wright CS, Hovatta O, Margara R, Trew G, Winston RM, Franks S & Hardy K 1999 Effects of follicle-stimulating hormone and serum substitution on the in-vitro growth of human ovarian follicles. Human Reproduction 1555–1562. (https://doi.org/10.1093/humrep/14.6.1555)
Xiao S, Zhang J, Romero MM, Smith KN, Shea LD & Woodruff TK 2015 In vitro follicle growth supports human oocyte meiotic maturation. Scientific Reports 17323 . (https://doi.org/10.1038/srep17323)
Xu M, Barrett SL, West-Farrell E, Kondapalli LA, Kiesewetter SE, Shea LD & Woodruff TK 2009 In vitro grown human ovarian follicles from cancer patients support oocyte growth. Human Reproduction 2531–2540. (https://doi.org/10.1093/humrep/dep228)
Zhang H & Liu K 2015 Cellular and molecular regulation of the activation of mammalian primordial follicles: somatic cells initiate follicle activation in adulthood. Human Reproduction Update 779–786. (https://doi.org/10.1093/humupd/dmv037)
