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Reproduction and Early Development Research Group, The Light Laboratories, Leeds Institute of Genetics, Health and Therapeutics, University of Leeds, Clarendon Way, Leeds LS2 9JT, UK
Correspondence should be addressed to H M Picton; Email: h.m.picton{at}leeds.ac.uk
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Abstract |
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 Top Abstract IntroductionOverview of follicle and...Follicle and oocyte growth...The follicle culture environment...Methods for harvesting follicles...Culture systems for the...The IVM of oocytesFuture prospects for the...ConclusionDeclaration of interestReferences |
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Introduction |
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TopAbstractIntroductionOverview of follicle and...Follicle and oocyte growth...The follicle culture environment...Methods for harvesting follicles...Culture systems for the...The IVM of oocytesFuture prospects for the...ConclusionDeclaration of interestReferences |
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Overview of follicle and oocyte development in vivo |
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TopAbstractIntroductionOverview of follicle and...Follicle and oocyte growth...The follicle culture environment...Methods for harvesting follicles...Culture systems for the...The IVM of oocytesFuture prospects for the...ConclusionDeclaration of interestReferences |
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Recent research has revealed that oocyte secreted factors regulate the initiation of primordial follicle growth (see Picton 2001 for review) and moderate the known trophic actions of the gonadotrophins FSH and LH on preantral and antral follicle growth (Knight & Glister 2006, McNatty et al. 2007, Webb & Campbell 2007). A plethora of putative regulators of folliculogenesis in vivo have been identified, some examples of these factors are shown in Fig. 1. The tyrosine kinase receptor Kit and the two different isoforms of its ligand, kit ligand (KL), have been localised to oocytes and GCs and have been shown to promote oocyte growth and maintenance of meiotic arrest in response to FSH receptor (FSHR) levels. While low concentrations of FSH promote oocyte growth by increasing KL-2 expression and by reducing the ratio of KL-1/KL-2, high concentrations of FSH enhance follicle development but impair oocyte growth (Thomas & Vanderhyden 2007). Other regulators of follicle growth include epidermal growth factor (EGF; Qu et al. 2000) and its receptor; activin (Hulshof et al. 1997, Telfer et al. 2008); basic fibroblastic growth factor (bFGF; Shikone et al. 1992); members of the insulin like growth factor (IGF) family and their binding proteins (Thomas et al. 2007), transforming growth factor-β (TGFB) superfamily members (Knight & Glister 2006) including somatic derived anti-Müllerian hormone (AMH), oocyte derived growth differentiation factor-9 (GDF9; McGrath et al. 1995, Dong et al. 1996); and the bone morphogenetic proteins (BMPs) especially BMP4, BMP7 and BMP15 (Shimasaki et al. 1999, Otsuka et al. 2001). Other factors such as retinoblastoma protein (RB1) may also be important (Bukovsky et al. 1995). Changes in follicle morphology and cell number, together with the stage specific follicular responsiveness to the growth factors detailed here and the development of steroidogenic capacity can be used as functional markers to confirm the normality of follicle development in vitro.
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Follicle and oocyte growth and development in vitro |
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TopAbstractIntroductionOverview of follicle and...Follicle and oocyte growth...The follicle culture environment...Methods for harvesting follicles...Culture systems for the...The IVM of oocytesFuture prospects for the...ConclusionDeclaration of interestReferences |
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The follicle culture environment in vitro |
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TopAbstractIntroductionOverview of follicle and...Follicle and oocyte growth...The follicle culture environment...Methods for harvesting follicles...Culture systems for the...The IVM of oocytesFuture prospects for the...ConclusionDeclaration of interestReferences |
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An important consideration when developing a culture medium for the IVG and maturation of follicles is the type and concentration of energy substrate included in the base medium. Basic research has shown that glucose is a common energy source for almost all types of animal cells; however, over the last few decades, it has become clear that mature oocytes of many mammalian species, including mice (Biggers et al. 1967, Leese & Barton 1984, Harris et al. 2007, 2008), cows (Gandolfi et al. 1998) and humans (Roberts et al. 2002, Harris & Picton 2007) preferentially metabolise pyruvate and have little capacity for the metabolism of glucose. Laboratory research has also shown that mouse oocytes have a high capacity for the consumption of pyruvate and oxygen at all stages of follicle development (Harris et al. 2008) while glucose consumption is below detection levels (Biggers et al. 1967, Eppig 1976, Harris et al. 2007). As a consequence of this evidence, glucose would be preferentially utilised by the somatic cells as the primary energy substrate in IVG media supporting follicle growth, whereas IVM media should be supplemented with pyruvate to support the energy needs of the maturing oocyte.
The inclusion of serum from homologous or heterologous sources in IVG and IVM media has ignited considerable debate. Serum contains proteins, amino acids, carbohydrates, trace elements, hormones, growth factors and extracellular matrix components which promote cell adhesion and proliferation. Extracellular matrix has been shown to moderate follicle survival in organ culture (Hovatta et al. 1999, Scott et al. 2004a). Serum acts as a source of albumin, which balances the osmolality and scavenges potentially harmful molecules and metal ions that can act as a source of free oxygen radicals. Serum also acts as a source of precursors for steroid biosynthesis (Hulshof et al. 1995). However, the inclusion of serum frequently blunts the response of proliferating cells to physiological doses of growth factors and it can act as a source of transmissible pathogens and viruses such as hepatitis B and C and bovine viral diarrhoea virus. Finally, the addition of serum in long-term cultures has been shown to lead to epigenetic modifications and foetal overgrowth syndromes (for example see Young et al. 1998, van Wagtendonk-de Leeuw et al. 2000). If serum is omitted from culture media, it is necessary to supplement the media with purified serum albumen or synthetic serum substitutes as a carrier molecule. Comparisons of the efficacy of IVG serum-based versus serum-free culture systems have been conducted in mice. Eppig et al. (1998) compared oocyte-GC complex development in culture medium supplemented with either foetal calf serum (FCS) or bovine serum albumin (BSA). More oocytes cultured with FCS cleaved after fertilisation and were competent to develop to the blastocyst stage than those cultured in BSA alone. In contrast, we and other authors have shown that FCS increased oocyte extrusion and follicle degeneration in isolated ovine follicles (for example Newton et al. 1999). Similarly, the replacement of FCS with BSA increased the percentage of morphologically intact preantral cat follicles by up to 30% (Jewgenow 1998).
There is a clear need to optimise the pH, temperature and oxygen tension of the culture environment in order to maximise the potential of in vitro derived oocytes (Ye et al. 2007). For example, although murine ovarian tissue is exposed to plasma oxygen levels of about 5% in vivo, mouse follicles grown in vitro in 5% oxygen had a higher frequency of mature oocytes with incorrectly aligned chromosomes and many died prematurely when compared with follicles cultured in 20% oxygen (Hu et al. 2001). Similarly, sheep follicles grown in 5% oxygen had reduced antral cavity formation and increased lactate and glucose consumption when grown under hypoxic conditions of 5% oxygen relative to sized matched counterparts that were grown in 20% oxygen (Jin et al. 2007). Additionally, the temperature of the media used for tissue transportation and holding prior to follicle harvest must be optimised to minimise apoptosis (Schmidt et al. 2003, Lucci et al. 2004).
Follicle culture practitioners need to consider the direct effects of long-term culture and sub-optimal culture environments on the epigenetic information and methylation status of key imprinted genes in oocytes (Huntriss & Picton 2008). This issue is of particular relevance to IVG and maturation strategies as epigenetic information that is essential for normal development is acquired during gametogenesis and a growing body of evidence suggests that assisted reproductive technologies, and especially those involving extended culture and invasive micromanipulations may affect these complex processes and lead to disorders of genomic imprinting (Huntriss & Picton 2008). Imprinted genes including the maternally expressed H19 gene, IGF2 and also epigenetic regulators such as members of the DNA methyltransferases family DNMT1 and DNMT3A have been shown to be aberrantly expressed following assisted reproduction technologies (Fernandez-Gonzalez et al. 2004, Sagirkaya et al. 2007). Analysis of the DNA methylation status of differentially methylated regions of the imprinted genes H19, paternally expressed gene 1/mesoderm-specific transcript (Peg1/Mest) and Igf2r in fully grown murine, germinal vesicle- staged oocytes derived in vitro from preantral follicles compared with in vivo grown counterparts indicated a loss of methylation at the Igf2r and Mest/Peg1 loci, as well as a gain of methylation of H19 (Kerjean et al. 2003). Furthermore, demethylation of the Peg1/Mest gene differentially methylated region has been observed in murine oocytes after extended culture (Imamura et al. 2005) suggesting that maternal imprints may be susceptible to demethylation during follicle culture. Further confirmation of this observation is provided following the detection of abnormal hypermethylation at the H19 locus in human oocytes matured in vitro (Borghol et al. 2006). Culture induced epigenetics disruption has been associated with the additions to FCS to different media (Khosla et al. 2001, Fernandez-Gonzalez et al. 2004). Although the exact mechanisms of imprinting disruption following extended culture are unclear, it is likely that cellular stress induced by suboptimal culture environments is highly relevant with the latter being manifest by the accumulation of NH3 in spent culture media (Lane & Gardner 2003). It is, however, important to note that there is a high degree of discordance between the imprinting status of genes in humans and other species (Morison et al. 2005) and there is variation in the regulation of epigenetic information including DNA methylation during preimplantation development between mammalian species (Beaujean et al. 2004, Fulka et al. 2004, Vassena et al. 2005). Consequently, it is vital that epigenetic health checks are conducted on all oocytes derived following IVG and IVM before these tissues can be used in a clinical setting.
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Methods for harvesting follicles for culture |
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TopAbstractIntroductionOverview of follicle and...Follicle and oocyte growth...The follicle culture environment...Methods for harvesting follicles...Culture systems for the...The IVM of oocytesFuture prospects for the...ConclusionDeclaration of interestReferences |
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Culture systems for the IVG and maturation of mammalian follicles and oocytes |
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TopAbstractIntroductionOverview of follicle and...Follicle and oocyte growth...The follicle culture environment...Methods for harvesting follicles...Culture systems for the...The IVM of oocytesFuture prospects for the...ConclusionDeclaration of interestReferences |
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Primordial and primary follicle culture
To maximise the reproductive potential of fresh and cryopreserved ovarian tissue it is necessary to develop culture strategies that support the activation and sustained IVG of primordial follicles as this is the most abundant stage of follicles development present in mammalian ovaries. Initial attempts at primordial follicle growth in vitro focussed on the growth of isolated follicles. While the data clearly show that it is possible to harvest rodent primordial follicles by mechanical means, in the tougher cortex of large animals and humans primordial follicles have to be isolated using enzymatic methods (Greenwald & Moor 1989; and human Oktay et al. 1997, Abir et al. 1999, Huntriss et al. 2002, Rice et al. 2008). Consequently, although a high percentage of primordial follicles are viable following extraction (
71%: Oktay et al. 1997), and growth can be supported for 24 h within a collagen gel (Abir et al. 1999), beyond this time follicles rapidly lose their three dimensional structure (Spears et al. 1994, Cortvrindt et al. 1996), the follicles collapse, the pregranulosa cells migrate away from the oocyte and/or oocyte extrusion and degeneration occurs. It is possible that the lack of influence from the extra-follicular cortical cells contributes to the demise of isolated primordial follicles in vitro as the presence of stromal cells around primordial follicles has been shown to improve initial culture success (Osborn et al. 1997). In contrast to the majority of publications, a recent report demonstrated that lectin-aggregated, serum-free cultures supported the long-term survival and growth of isolated ovine primordial follicles from both fresh and cryopreserved neonatal lamb ovary ovarian cortex (Muruvi et al. 2005). Oocyte growth and GC differentiation were further enhanced in lectin-aggregated cultures following exposure to KL in vitro and the oocytes reached diameters equivalent to those of oocytes from primary follicles derived in vivo (Muruvi et al. 2005). While these recent results are encouraging, they are by no means definitive as the optimal methodology for primordial follicle growth in vitro.
An alternative approach that has consistently proved successful for primordial follicle culture in a number of species is to grow these early stages in situ within fragments of the ovarian cortex. In early in situ culture experiments, large numbers of oocytes underwent atresia (Blandau et al. 1965), and no follicular growth was observed (Baker & Neal 1974). However, Eppig & O'Brien (1996) reported the birth of live pups from oocytes derived from primordial follicles grown in situ following the culture of foetal mouse ovaries. Two subsequent studies have also produced live offspring following the growth of oocytes present within cryopreserved ovaries (Liu et al. 2001) and foetal ovaries obtained at E12.5 (Obata et al. 2002). In situ cultures of primordial and primary follicles can be conducted by culturing fragments of ovarian cortex directly or by grafting the cortical fragments beneath the highly vascularised chorioallantoic membrane of chick embryos (Fortune et al. 2000, Cushman et al. 2002, Qureshi et al. 2008). Whichever route is adopted, culture of thin cortical strips: i) avoids the damage caused by the follicle harvesting procedure; ii) minimises the chances of necrosis by maximising the surface area of the tissue for gaseous exchange and fulfilment of the nutrient requirements; and iii) provides a complex support system that more closely resembles the ovary in vivo as the follicles remain both intact and in contact with the surrounding stromal cells, which provide the local biochemical control pathways that trigger the initiation of follicle growth.
In situ cortical cultures have been conducted for a number of species over varying time frames with or without extracellular matrix and high levels of serum (Hovatta et al. 1999, Wright et al. 1999) or in serum-free medium (Picton et al. 1999b, Webber et al. 2007, Telfer et al. 2008). During the in situ culture pieces of ovarian cortex are placed onto a physical support, such as a porous membrane insert, in culture wells and the tissue is then covered with a meniscus of culture media (Devine et al. 2002). Using this approach, follicles may remain viable and are morphologically normal after 1–3 weeks of culture. Using this method, recruitment of primordial follicle growth has been demonstrated in a number of species (cows: Wandji et al. 1996, 1997, Braw-Tal & Yossefi 1997, Fortune et al. 1998; baboons: Fortune et al. 1998, hamsters: Yu & Roy 1999, humans: Hovatta et al. 1999, Picton et al. 1999b, Wright et al. 1999). Furthermore, multilayer preantral follicles have been isolated following the culture of fragments of ovarian cortex (Telfer et al. 2008) or after culture of whole mouse ovaries (Eppig & O'Brien 1996). Despite these successes, not all cortical strip culture systems are equally effective and further optimisation of the oxygen tension and growth additives are required to sustain high levels of primordial and primary follicle growth in vitro.
The problems encountered during the culture of isolated primordial follicles are replicated during primary follicle IVG. However, recent data have shown that it is possible to use an attachment based culture system to drive secondary follicles in vitro from isolated primary follicles harvested from fresh or cryopreserved neonatal lamb ovaries (Muruvi et al. 2008). Here, the oocytes from isolated primary follicles were shown to be able to reassemble functional ‘follicle units’ in vitro and to support the sustained growth of these reorganised follicles to secondary stages over more than 42 days in vitro. After 7, 11 and 30 days of culture increases in follicle diameter of 7 µm (Nuttinck et al. 1993), 19 µm (Itoh & Hoshi 2000) and 35 µm (Saha et al. 2000) respectively, have been reported for small bovine follicles. In this cohort of studies, follicle survival and growth depended on the use of feeder layers of somatic cells.
Preantral-antral follicle culture
Unlike primordial and primary follicle culture, pre-antral–antral cultures are rarely conduced in situ but rather follicle isolation appears to be crucial for future development in vitro. The growth characteristics of preantral follicles in vitro and the duration of culture are highly dependent on the culture system and the species of interest. Preantral follicles have been cultured with serum on a plastic substratum (Figueiredo et al. 1994, Jewgenow 1998), grown on somatic cell monolayers (Itoh & Hoshi 2000), or in serum-free environments (Newton et al. 1999, Gutierrez et al. 2000). Preantral follicle culture systems broadly fall into two categories: i) culture systems that allow follicle attachment and loss of follicle integrity as illustrated in Fig. 3 (mouse: Eppig & Schroeder 1989, Cortvrindt et al. 1996); or ii) three dimensional (3-D) culture systems that maintain follicle integrity. Three dimensional systems have been achieved through the use of physiological serum-free media approaches as shown in Fig. 4 (sheep: Newton et al. 1999; human: Picton et al. 1999b, Telfer et al. 2008), through the use of hydrophobic membranes that prevent attachment (mouse: Nayudu & Osborn 1992); or via encapsulation in collagen (mouse: Carroll et al. 1991a; pig: Hirao et al. 1994) or alginate gels (Xu et al. 2006, West et al. 2007).
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Three dimensional IVG systems originated with the culture of murine preantral follicles (for example: Eppig & Schroeder 1989, Carroll et al. 1991b, Nayudu & Osborn 1992, Hartshorne et al. 1994, Spears et al. 1994, Cortvrindt et al. 1996). Using this approach, intact follicles are cultured on hydrophobic membranes that inhibit cellular attachment and migration (Nayudu & Osborn 1992). Alternatively, mouse follicles are cultured in microdrops under mineral oil and grown from mid-preantral sized follicles to the Graafian stage in about 6 days. The daily transfer of follicles to a new culture well prevents follicular attachment (Boland et al. 1993). The antral follicles derived using the three-dimensional system have normal morphology and steroid production (Nayudu & Osborn 1992, Hartshorne et al. 1994, Spears et al. 1994); furthermore, up to 80% of these follicles ‘ovulate’ in vitro in response to an exogenous ovulatory stimulus (Rose et al. 1999) and the oocytes are healthy and can be fertilised with the production of live young (Spears et al. 1994).
A number of different approaches for three-dimensional culture have been tested in mice; these methods may be more compatible with the IVG requirements of the larger follicles collected from domestic ruminants and humans. Anchorage independent cultures of whole murine follicles have utilised inverted drop cultures of preantral follicles of 180–240 µm diameter (Wycherley et al. 2004). Rotating wall suspension cultures have also been tried (Rowghani et al. 2004). Alternatively, encapsulation of follicles in inert alginate hydrogels has been shown to be a good option for the sustained growth of larger follicles. Alginate is a linear polysaccharide copolymer of β-D-mannuronic acid and 
-L-guluronic acid, which is produced by brown algae (Haug et al. 1967). It is thought that the alginate matrix mimics the stromal microenvironment of the ovary and as such will better support the growth and maturation of multilayered secondary follicles in large animals, primates and humans (Kreeger et al. 2007). Early reports demonstrated that murine follicles grown in alginate capsules reached the sizes typically observed in vivo (Pangas et al. 2003, Kreeger et al. 2007). Extended growth, coordinated differentiation of follicular cell types, antral cavity formation, theca cell differentiation, and oocyte maturation and hormone production were achieved by decreasing matrix stiffness and by altering the alginate scaffold concentration (Xu et al. 2006, West et al. 2007). This method of in vitro folliculogenesis from preantral stages has also resulted in the production of live young (Xu et al. 2006). While these data are very encouraging, it is important to note that the follicle and oocyte growth rates in all of these reports are increased over in vivo grown counterparts. The health of oocytes derived from follicles with accelerated growth rates needs to be fully verified before these systems can be considered suitable for the clinical use of in vitro derived oocytes.
Preantral follicle culture in large domestic animals and humans
Despite the many advances in rodent follicle culture detailed above, these achievements have proved difficult to replicate in large animals and humans. One of the main reasons for the lack of progress is that it is technically more difficult to recover follicles from these species, as the cortex has a more dense and fibrous consistency (Telfer 1996) and follicle isolation frequently requires the use of aggressive enzyme-based protocols. Nevertheless, in recent years, it has proved possible to maintain the three-dimensional integrity of ruminant (Newton et al. 1999, Gutierrez et al. 2000, Picton et al. 2003, Thomas et al. 2007) and human follicles (Picton et al. 1999b, Telfer et al. 2008). The philosophy that underpins this system is that cultured GCs form distinct clumps of rounded cells that closely resemble the morphology (Chang et al. 1977) and steroidogenic phentoype of GCs in vivo (Campbell et al. 1996, Picton et al. 1999a) rather than acquiring a fibroblastic character associated with attachment to the plastic culture dish. This species–specific system maintains follicle integrity (Fig. 4) and follicle cells retain their normal ultrastructure (Jin et al. 2004). Antral cavity formation occurs in 50% of isolated ovine secondary follicles with a start size of 180–220µm after 12–14 days of culture, with growth being supported up to diameters of 1.2 mm over 30 days in both fresh and cryopreserved tissues (Newton et al. 1999). Using this system, follicle growth rates can be manipulated by the presence/absence of the theca cell layer, as well as the presence/absence of basement membrane components in the culture media. These data support the concept first observed in murine follicles by Boland & Gosden (1994), that the theca cells have biochemical effects on oocyte growth and development in vitro, which are mediated via the GCs (Kotsuji et al. 1994, Richard & Sirard 1996). Additionally, on the provision of appropriate concentrations of androgen substrate, sheep follicles grown over 30 days resulted in the biosynthesis of limited, but physiological levels of oestradiol in vitro (Newton et al. 1999). Although the follicular growth rates recorded in this system in vitro are slightly accelerated compared with growth rates in adult animals, they closely mimic the growth rates of similar staged follicles in foetal ovaries in vivo. This extended growth phase is important as it facilitates the sequential accumulation of the payload of RNAs and proteins required to support oocyte developmental competence.
The potential of three-dimensional pre-antral follicle culture has been demonstrated in other domestic species including pigs (Wu et al. 2001); cattle (Gutierrez et al. 2000, McCaffery et al. 2000, Thomas et al. 2001, 2007) and humans (Picton et al. 1999b, Telfer et al. 2008). The IVG of porcine follicles has produced a small number of meiotically competent oocytes following growth in both collagen gels (Hirao et al. 1994), and in uncoated culture wells (Telfer et al. 2000). In vitro fertilisation of IVG-derived porcine metaphase II oocytes resulted in limited blastocyst production (Wu et al. 2001). The IVG of human pre-antral graulosa-oocyte-complexes and whole follicles have resulted in development up to antral sizes following enzymatic isolation and culture within agar (Roy & Treacy 1993) or collagen (Abir et al. 1999) gels. Mechanical isolation and IVG have also been demonstrated (Abir et al. 1997), although levels of atresia following culture were found to be high. In the most recent report (Telfer et al. 2008), pre-antral follicles have been isolated mechanically from cortical tissue after 6 days of culture and antral cavity formation occurred in the isolated follicles very rapidly, within 2 days of isolation. The meiotic competence and developmental capacity of human oocytes grown from pre-antral stages in vitro has yet to be confirmed.
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The IVM of oocytes |
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TopAbstractIntroductionOverview of follicle and...Follicle and oocyte growth...The follicle culture environment...Methods for harvesting follicles...Culture systems for the...The IVM of oocytesFuture prospects for the...ConclusionDeclaration of interestReferences |
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Future prospects for the IVG and maturation of follicles |
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TopAbstractIntroductionOverview of follicle and...Follicle and oocyte growth...The follicle culture environment...Methods for harvesting follicles...Culture systems for the...The IVM of oocytesFuture prospects for the...ConclusionDeclaration of interestReferences |
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Conclusion |
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TopAbstractIntroductionOverview of follicle and...Follicle and oocyte growth...The follicle culture environment...Methods for harvesting follicles...Culture systems for the...The IVM of oocytesFuture prospects for the...ConclusionDeclaration of interestReferences |
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Declaration of interest |
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TopAbstractIntroductionOverview of follicle and...Follicle and oocyte growth...The follicle culture environment...Methods for harvesting follicles...Culture systems for the...The IVM of oocytesFuture prospects for the...ConclusionDeclaration of interestReferences |
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Funding |
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Received July 5, 2008
First decision August 4, 2008
Revised manuscript received September 15, 2008
Accepted October 7, 2008
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References |
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TopAbstractIntroductionOverview of follicle and...Follicle and oocyte growth...The follicle culture environment...Methods for harvesting follicles...Culture systems for the...The IVM of oocytesFuture prospects for the...ConclusionDeclaration of interestReferences |
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