Abstract
With the advent of culture models for studying the process of ovarian folliculogenesis some 40 years ago, opportunities arose for the more systematic evaluation of the factors that regulated ovarian function [1]. The initial focus of studies using cultured follicles emphasized two of the then widely recognized roles of the follicle in mammals: the production of ovarian steroid hormones and of viable oocytes during the process of ovulation. As our understanding of the molecular and cellular complexity of this tissue compartment has evolved and deepened, so too has the need to redefine the major functions of the follicle at both local ovarian and systemic levels in the context of reproduction in mammalian species, especially as it relates to the origins and treatments for human infertility [2]. Thus, a shift in the motivation to use cultured follicles in humans has taken place owing primarily to the rapidly evolving field of fertility preservation.
Introduction
In vitro culture systems for ovarian follicles are enabling tools for advancing the study of folliculogenesis and the development of fertility preservation techniques. Folliculogenesis is a complex process regulated by endocrine, paracrine, autocrine and juxtacrine factors, as well as cell-matrix interactions, and can be difficult to study in vivo. In vitro culture systems provide a controlled environment in which to investigate the mechanisms driving follicle development. These systems have enabled significant discoveries about the influence of hormones, cell-cell, and cell-matrix interactions in folliculogenesis [1]. In addition to these fundamental observations, these culture systems are providing a foundation for the development of systems for fertility preservation and restoration for cancer survivors and other women with complex medical conditions, such as inflammatory and autoimmune diseases [2–4]. The increase in survival rates for young women with cancer has prompted the need for fertility preservation techniques [5]. Life-saving cancer treatments, such as chemotherapy and radiation, threaten fertility by diminishing the immature follicle pool and triggering early menopause. For patients unable to cryopreserve oocytes or embryos, the only treatment currently available is cryopreservation and subsequent autotransplantation of ovarian tissue, which incurs the risk of reintroducing cancer cells into the patient [6]. The successful development of follicle culture systems could circumvent this risk, by allowing in vitro follicle maturation to obtain fertilizable oocytes from immature follicles, followed by in vitro fertilization and embryo transfer. Follicle culture systems have had some success; however, further developments are necessary to achieve the consistent growth of human follicles to produce fertilizable, developmentally competent oocytes [7, 8], especially when starting from the earliest stage follicles.
Stromal tissue is essential for the development of the earliest stage follicles. The ovarian stroma contains interstitial–theca cells, neurons, blood vessels, and macrophages. Secondary follicles are routinely grown individually using in vitro culture systems; however, primordial and primary follicles depend on distinct microenvironments in order to survive in culture. Primordial follicles require a rigid microenvironment [9] and primary follicles are not competent to grow individually and instead require signaling from other cells, including the stroma [10–12]. These early stage follicles typically grow best in organ culture-containing stromal components [13–19]. Hence, the stroma has a clear influence on follicle development. Stromal cells provide structural support and have complex bidirectional paracrine signaling with the follicle [20–23]. Moreover, it is widely hypothesized that stromal cells are recruited by the follicle and differentiate into theca cells [24]. Nevertheless, the specific mechanisms of action remain unclear. In vitro culture systems that contain stromal cells or stromal cell equivalents (e.g., fibroblasts) may be an effective tool for investigating the mechanisms by which stromal cells activate the early stage follicles and may ultimately be translated toward strategies for fertility preservation for cancer patients.
In this chapter, we discuss ovarian stromal cells and in vitro follicle culture systems. Integrating stromal cells into current follicle culture systems will better simulate the natural ovarian microenvironment and could lead to the elucidation of the mechanisms by which these follicles are activated. In addition, stromal cell co-culture could improve early follicle growth and survival, which are essential for the successful translation of in vitro culture systems to primate and human follicles. The current knowledge base for the culture of somatic and stromal cells is discussed, with an emphasis on three-dimensional culture systems using biomaterials. In addition, the potential and challenges of co-culture is discussed by drawing from recent work with co-culture systems in tissue engineering.
Roles of Ovarian Stromal Cells in Follicle Development
Folliculogenesis is the process by which primordial follicles develop to antral follicles that ovulate fertilizable oocytes (Figure 30.1). The follicle is the functional unit of the ovary and is composed of a germ cell (the oocyte) and layers of somatic cells (granulosa and theca cells). Primordial follicles are the most immature class and are found during embryonic (human) or immediately post-embryonic (murine) life. Primordial follicles are the most abundant follicle class and represent the ovarian reserve, or the potential of the organ to continuously provide systemic hormone support and fertilizable eggs in a monthly cycle. The oocyte within primordial follicles is arrested in the first meiotic prophase and is surrounded by a single layer of squamous granulosa cells. Follicle activation from the non-replenishable ovarian reserve is a process that is not well understood. Cohorts of primordial follicles are activated and form primary follicles, which have larger oocytes and a layer of cuboidal granulosa cells enclosed by a basement membrane. Subsequently, the granulosa cells proliferate and form several layers around the oocyte. At the same time, theca cells begin to surround the basement membrane of the follicle. These follicles are called secondary and multilayer follicles. Under the influence of follicle stimulating hormone (FSH), the granulosa cells proliferate and differentiate into cumulus (surrounding the oocyte) and mural (inside of the basement membrane) granulosa cells. Likewise, the theca cells differentiate into theca interna (androgen secreting cells) and externa (connective and supportive tissue). The follicle increases in size and develops a fluid-filled cavity called an antrum. Under the influence of luteinizing hormone (LH), the oocytes in antral follicles complete the first meiotic cell division and then pause in the second meiotic metaphase. These oocytes are capable of fertilization. The overall goal of follicle culture systems is to reproduce this entire process in vitro.
Figure 30.1 Folliculogenesis. Primordial follicles develop to antral follicles, which are capable of producing fertilizable oocytes. The follicle microenvironment, which includes ovarian stromal cells, is hypothesized to have significant roles in the activation of primordial follicles and the recruitment/differentiation of theca cells. Most follicle culture systems focus on secondary or multilayer follicles and produce antral follicles with fertilizable oocytes. Primordial and primary follicles only activate or mature in vitro when cultured under conditions that mimic their physiologic microenvironment. The culture of these follicles is often performed as an organ culture.
Ovarian stromal cells may have significant roles in folliculogenesis, particularly in the activation of primordial follicles and the differentiation of theca cells. Stromal cells, which are similar in morphology to fibroblasts, make up the connective tissue throughout the ovary and around follicles. Stromal cells are assumed to arise from a population of unspecialized mesenchymal stem cells [25]. The morphology of the stromal tissue varies between the cortex and medulla of the ovary [26]. In the cortex, the stromal cells are organized parallel to the surface and have a rounded structure. In the medulla, the cells exhibit random organization and have an elongated structure, and they are often referred to as interstitial or luteinized cells. The stromal cells in the medulla are believed to be further differentiated toward theca cells and have greater steroidogenic capacity than the stromal cells in the cortex [27]. Stromal cells are often identified as mesenchymal cells that lack theca cell markers, such as the LH receptor (LHR), steroidogenic acute regulatory protein (StAR), 3β-hydroxysteroid dehydrogenase (3β-HSD), and 17α-hydroxylase (CYP17A1) [27, 28]. Chicken ovalbumin upstream promoter-transcription factor (COUP-TFII) has been used to stain stromal cells, but it is also expressed in theca cells [29]. Recently, Laronda et al. proposed high mobility group box 1 protein (HMGB1) as a stromal cell marker [30]. HMGB1 is expressed in ovarian stromal cells and oocyte nuclei, but not in granulosa or theca cells, making it a potentially useful marker to distinguish somatic cells. Stromal cells signal via paracrine factors and influence early follicle development. For example, bone morphogenetic proteins 4 and 7 (BMP-4 and -7), secreted by stromal and/or theca cells have been identified as positive regulators of the primordial-to-primary follicle transition [31, 32]. Moreover, it is widely hypothesized that stromal cells are recruited by the follicle via paracrine factors and differentiate into theca cells. This hypothesis was first proposed by Dubreuil in 1946 [33] and later reiterated by others [24, 34]. To date, a number of paracrine factors that regulate theca cell recruitment and differentiation have been identified, such as insulin-like growth factor (IGF-1), kit ligand (KL), and basic fibroblastic growth factor (bFGF) [23, 35, 36]. Nevertheless, the conclusive evidence for this hypothesis remains elusive. In vitro culture systems of follicles and stromal cells may be an enabling tool to elucidate the functions of the ovarian stroma, signaling mechanisms, and roles in follicle development.
Studies Investigating the Roles of the Ovarian Stroma
Follicle-to-stroma Paracrine Signaling
Early stromal cell experiments established paracrine signaling from the follicle to the stroma. In 1995, Magoffin and Magarelli demonstrated that granulosa cells of developing follicles secrete a signal that stimulates theca cell differentiation [20]. In this experiment, isolated theca–interstitial cells from rat ovaries were cultured in media conditioned by follicles and assayed for androgen secretion. The conditioned media was found to stimulate androgen secretion, which suggested the presence of a theca differentiation factor secreted by the follicle. Later, Magarelli et al. identified an increase in the mRNA expression of the LHR and various theca steroidogenic enzymes, including cholesterol side-chain cleavage (P450scc), 3β-HSD, and CYP17A1 in response to this conditioned media [22]. Also, Magarelli et al. showed that this differentiation signal was both gonadotropin (FSH) and developmentally regulated, as only pre-antral follicles with 2–5 layers of granulosa cells produced the signal [22]. To date, the identity of this signal has not yet been purified; however, several proteins in the range of 19–24 kD that act synergistically may be responsible for this action [21]. In a series of candidate signaling studies, the combination of two granulosa produced peptides, IGF-I and KL, were found to increase androgen production and gene expression of androgenic factors in rat theca–interstitial cells [23]. Other potential regulating factors include growth differentiation factor-9 (GDF-9), activin, inhibin, and follistatin [21, 37, 38]. In sum, these experiments support the notion that follicle-to-stroma signaling exists.
Additional studies have demonstrated the effects of follicle-to-stroma signaling on theca cell recruitment, stromal cell proliferation, and the primordial-to-primary follicle transition. Parott and Skinner treated ovary fragments and isolated stromal–interstitial cells with KL, which was hypothesized to be a “theca cell organizer” secreted by granulosa cells [35]. KL was found to significantly increase the percentage of theca cell layer thickness of primary follicles in organ culture. This result suggests KL helps to recruit theca cells from the stroma. Furthermore, KL was found to stimulate ovarian stromal cell proliferation in a dose-dependent manner. Treatment with KL did not affect stromal cell androstenedione or progesterone production. Hence, KL did not promote theca cell differentiation, which is consistent with research that has determined that theca cell differentiation is controlled by a synergy of multiple factors [21–23]. Later, Nilsson and Skinner identified the roles of KL and bFGF in the primordial-to-primary follicle transition [36]. The combination of these factors decreased the percentage of primordial follicles and increased the percentage of primary, pre-antral and antral follicles. Overall, these studies build upon the earlier work by identifying the possible roles of stromal cells in folliculogenesis.
To further explore the effects of paracrine signaling, stromal cells have been co-cultured with granulosa and theca cells. Orisaka et al. developed a co-culture system that separates two cell populations via a collagen membrane, which permits the diffusion of factors smaller than 12.5 kD [27]. In these studies, bovine stromal cells from the cortex and medulla of the ovary were cultured with and without granulosa cells. Co-culture with granulosa cells increased the number of secreted lipid droplets, filopodia and mitochondria. Co-culture also stimulated androgen secretion in both cortex and medulla stromal cells. However, an increase in LH receptor mRNA was only observed in cortex stromal cells. Surprisingly, no increase in other theca markers was detected. While these data suggests theca cell differentiation, the evidence is not conclusive. No changes in the co-cultured granulosa cells were reported. Nevertheless, these experiments successfully demonstrated paracrine signaling from granulosa cells to the stromal cells in a co-culture system.
Stroma-to-follicle Paracrine Signaling
In addition to paracrine signaling from the follicle to the stroma, signaling from the stroma to the follicle has been studied. The bone morphogenetic proteins BMP-4 and BMP-7, which are expressed by stromal and/or theca cells have been linked to the primordial-to-primary follicle transition. Lee et al. demonstrated that BMP-7 promotes the activation of primordial follicles in vivo [32, 39]. BMP-7 was injected into the ovarian bursa of rats, which produced a decrease in the number of primordial follicles and an increase in the number of primary, pre-antral and antral follicles. Nilsson et al. achieved similar results using in-vitro organ culture and BMP-4 [40]. Thus, these studies support the notion that stroma-to-follicle signaling exists.
Organ Culture and the Activation of Primordial Follicles
Some of the most significant stromal cell experiments have been ovary organ culture experiments (in situ). These experiments culture follicles within thin fragments of ovarian stromal tissue. In 1996, Eppig and O’Brien achieved complete oocyte development in vitro using oocytes from primordial follicles of newborn mice [14]. This result was accomplished using a two-step strategy in which the ovaries of newborn mice were grown in organ culture for 8 days and then oocyte–granulosa cell complexes were isolated from the ovaries and cultured individually for an additional 14 days. Using this strategy, primordial follicles developed to produce mature oocytes. These oocytes were then fertilized in vitro and the resulting embryos were implanted to produce live offspring. When cultured individually outside of stromal tissue, primordial follicles rapidly lose their three-dimensional structure, pre-granulosa cells migrate away from the oocyte and oocyte extrusion/degeneration occurs [41]. Follicle culture in ovary organ fragments provides a complex support system that closely resembles the in-vivo ovary environment. Follicles maintain contact with the supporting stromal cells, which provide the local biochemical control pathways that trigger follicle activation [42]. Eppig and O’Brien’s method was later improved and translated to follicles from larger animals, such as cow and sheep [15–17]. Similarly, Laronda et al. demonstrated that human primordial follicles, which did not survive in culture isolated from ovarian tissue, survived, differentiated, and grew when cultured within ovarian cortical strips [18]. Telfer et al. also used this method to activate primordial human follicles [13]. Primordial follicles were matured to secondary follicles in organ culture and then cultured individually to the early antral stage with the treatment of activin. These experiments clearly demonstrate the role of the physiologic follicle microenvironment in regulating folliculogenesis. Some characteristics of the physiologic microenvironment, such as its mechanical rigidity, structure, or bioactivity, can be mimicked through the use of engineered biomaterials, but the important functions of stromal cells in folliculogenesis justify the integration of stromal cells into the current follicle culture system in order to achieve the activation of individual primordial and primary follicles in vitro.
Three-Dimensional Culture Systems for Ovarian Follicles
Most follicle culture systems have focused on isolated secondary/multilayer follicles or oocyte–granulosa cell complexes. Primordial and primary follicles do not mature when cultured individually in vitro. The majority of work has been completed using rodents due to the low cost and relatively short-growth period compared to larger animals. Follicle culture systems can be divided into two approaches: two-dimensional (flat) (Figure 30.2A) and three-dimensional (spherical) (Figure 30.2B). For detailed reviews of follicle culture systems see the following references [1, 43]. In two-dimensional follicle culture systems, follicles are cultured on flat surfaces such as tissue culture plastic (polystyrene), collagen, or polylysine. The first successful culture system using isolated mouse follicles was developed by Eppig in 1977 [44]. This system was later improved and achieved the birth of live offspring [14, 45]. While two-dimensional culture systems have proved to be successful at producing mature oocytes in mice, this approach has proven difficult to replicate with follicles from large animals and humans. The unnatural geometry of two-dimensional culture disrupts cell–cell communication and causes the granulosa cells to break through the basement membrane, migrate away from the oocyte and attach to the two-dimensional surface [41, 46–48]. Thus, in order to culture larger follicles and achieve the goal of maturing human follicles in vitro, three-dimensional culture systems are necessary to properly support the developing follicle.
Three-dimensional follicle culture systems maintain the natural spherical geometry and cell–cell interactions of the follicle. Three-dimensional culture systems have utilized polylysine or collagen-coated substrates [47, 49], hydrophobic membranes [50], rotating walls and orbiting test tubes [51], mineral oil with daily follicle transfer [52], inverted culture [53], and serum-free media [13]. These systems have been successful at maintaining follicle geometry, preventing granulosa cell migration and minimizing attachment to the flat surface. Nevertheless, these systems still lack the ability to support large follicles for extended culture times. Fortunately, the application of biomaterials to follicle culture offers the potential to overcome these limitations and provide a true three-dimensional culture environment that mimics the physiologic ovary.
Figure 30.2 In vitro follicle culture systems. (A) In two-dimensional systems, follicles are cultured on flat surfaces such as tissue culture plastic (polystyrene). The unnatural geometry/mechanics of these systems disrupts cell–cell communication and causes the granulosa cells to break through the basement membrane, migrate away from the oocyte, and attach to the two-dimensional surface. Two-dimensional systems lack the ability to support large follicles for extended culture times. (B) In three-dimensional systems, follicles are cultured within biomaterial scaffolds, such as alginate. These systems maintain the natural spherical geometry and cell–cell interactions of the follicle.
Three-dimensional biomaterial scaffolds mimic in vivo cellular microenvironments better than flat two-dimensional surfaces [54]. The usage of biomaterial scaffolds has demonstrated the effect of geometry and mechanics on cell survival, proliferation, migration, gene expression and differentiation. For example, Bissell and coworkers demonstrated that human mammary epithelial cells display a spread phenotype when cultured on a two-dimensional surface, yet form normal acinar structures when cultured in a three-dimensional environment [55, 56]. Moreover, Tanaka et al. found that enhanced chondrogenesis resulted from the three-dimensional culture of embryonic stem cells compared to flat monolayer culture [57]. In addition to geometry, mechanics has a substantial effect on cell behavior. For example, Engler et al. identified that stem cell lineage was controlled, in part, by scaffold elasticity [58]. Mesenchymal stem cells were directed to neurogenic, myogenic and osteogenic lineages using soft, stiff and rigid matrices. Hence, these studies have established the need to provide the proper environment for cell culture. This task has been accomplished via the use of engineered biomaterials that permit the modification of physical properties. Thus, biomaterials have provided an effective strategy to mimic in vivo environments. Highly water-soluble polymer networks, called hydrogels, have been used as biomaterial scaffolds for follicle culture. Hydrogels support natural follicle growth by mimicking the microenvironment of the physiologic ovary. A variety of natural [7, 59–61] and synthetic [62] hydrogels have been used for three-dimensional follicle culture. Early hydrogels for follicle culture employed collagen [59, 63–67], which is an extracellular matrix protein that is prominent throughout the ovary [68]. Torrance et al. demonstrated the growth and survival of primary mouse follicles in collagen hydrogels, but did not achieve antrum formation [64]. Later, Hirao et al. and Joo et al. produced mature oocytes from pre-antral pig [67] and rat [59]follicles, respectively, and Sharma et al. achieved antral follicles from pre-antral buffalo follicles [65]. In addition to collagen, alginate, a common tissue engineering hydrogel, has shown great promise as a scaffold for follicle culture [7, 46, 69–75]. It is a naturally derived polysaccharide isolated from brown algae. Alginate is a block copolymer composed of blocks of (1–4)-linked β-D-mannuronic acid (M units) and its C-5 epimer α-L-guluronic acid (G units) [76]. These blocks can be either similar or alternating. Alginate chains can be crosslinked with divalent cations, such as Ca2+. As a result, alginate avoids the use of harmful chemicals, ultraviolet light or temperatures to crosslink the polymer network [1]. Crosslinking occurs via interaction of the carboxylic acid functional groups in the G-blocks. This crosslinking leads to the formation of a gel network while retaining cell viability and the cellular interactions within the follicle. Due to its hydrophilic nature, alginate discourages protein adsorption and cell attachment. Thus, cells are unable to specifically interact or bind with alginate. In order to overcome this limitation, alginate can be covalently linked to cell-adhesion molecules, such as RGD (Arg-Gly-Asp) peptide ligands, via its carboxylic acid functional group via carbodiimide chemistry [77]. In addition, alginate can be degraded with alginate lyase, an alginate specific enzyme, in order to safely remove the follicle from the hydrogel. Hence, due to its inert nature, gentle and rapid gelation and modularity, alginate remains a top candidate for follicle culture.
Figure 30.3 Stromal cell co-culture approaches. (A) Stromal cells can be cultured separately on a flat surface below the encapsulated follicle. This set up allows for paracrine signaling between the follicle and the stromal cells. (B) Stromal cells can be encapsulated inside the biomaterial scaffold with the follicle. This set up allows for paracrine signaling as well as cell–cell attachment and interaction with secreted extracellular matrix proteins.
Alginate is a successful encapsulating matrix for follicle culture. Experimentally, isolated follicles are suspended within drops of alginate, which are subsequently crosslinked in a Ca2+ solution. Each resulting alginate bead is then transferred to an individual well in a standard cell culture plate containing growth media. Alginate hydrogels have been shown to maintain the natural three-dimensional morphology of the developing follicle. This culture system has demonstrated the successful maturation of secondary to antral follicles with mice, goats, sheep, dogs, primate and human follicles [8, 72, 78–81]. Follicles grow, develop antrums and produce meiotically competent oocytes. These oocytes have been successfully fertilized in vitro and implanted into mice, which has yielded healthy, fertile offspring [74]. Follicle growth was found to be strongly influenced by the mechanical rigidity of the surrounding alginate matrix. Permissive environments (low concentrations of alginate) supported the most rapid follicle growth and highest rates of fertilization [1, 70, 71]. In addition, the incorporation of extracellular matrix (ECM) proteins, such as collagen, fibronectin and laminin, into the alginate matrix has been found to improve follicle growth and oocyte quality [46]. This culture system has also been used in combination with ovary organ culture to produce mature oocytes from primordial follicles [82]. Furthermore, alginate has been used to culture human follicles after cryopreservation [83]. Hence, alginate has proven to be powerful tool for the advancement of follicle culture.
Additional functionalities are being incorporated into the alginate system to enhance follicle development. A fibrin-alginate interpenetrating network (FA-IPN) was developed to provide dynamic cell-responsive mechanical properties to the culture system [75]. Fibrin, the natural polymer involved in blood clotting, and alginate are formulated simultaneously into a single hydrogel. As the follicle grows, proteases are secreted to degrade the fibrin, leaving only the non-degradable alginate matrix to support the follicle. Fibrin alone was not successful in promoting follicle development. This FA combination produces a more permissive environment than can be achieved with alginate alone. The rate of meiotically competent oocytes produced was 82%, which is significantly higher than alginate alone or any other reported in vitro culture system. By promoting interaction with the encapsulating matrix, degradable hydrogels better mimic the natural ovary microenvironment. Future follicle culture systems will likely build upon this example, leveraging more bioactive and biomimetic materials, in order to further improve oocyte quality and survival, which is essential for the complete translation to primate and human follicles.
Utility of Co-Culture for Ovarian Follicle Development
The next logical step in the development of follicle culture systems is the integration or co-culture of stromal cells (Figure 30.3). While biomaterials can be engineered to mimic the physical properties of the extracellular matrix, somatic cells must also be incorporated in order to fully recapitulate all the roles of the stromal compartment. As demonstrated via stromal cell and ovary organ culture experiments, stromal cells have a significant role in the activation of primordial/primary follicles and theca cell recruitment/differentiation. The paracrine signaling involved in these processes is hypothesized to be a complex time-dependent synergy of unidentified factors [21]. Cell–cell contact may also have an important role in these processes. For these reasons, the simple addition of candidate hormones and growth factors to the culture media has not yet achieved success. Until the precise spatial-temporal molecular signaling mechanisms that govern these processes are elucidated, stromal cells could be incorporated into follicle culture systems as a way to potentially culture earlier stage follicles and improve growth, survival and oocyte quality.
While only a few co-culture studies with stromal cells have been conducted, these experiments demonstrate positive results and motivate further investigation. Osborn et al. showed that the presence of stromal cells around isolated primordial follicles improved initial culture success [84]. Building upon this observation, Itoh and Hoshi co-cultured small pre-antral (primary and secondary) bovine follicles with ovarian mesenchymal cells, granulosa cells and skin fibroblasts for 30 days [85]. Compared to the non-co-culture controls, follicle viability was significantly increased in all three co-cultures (18.6, 17.1 and 10.0%, respectively) and follicle growth was significantly increased in the mesenchymal (15.4%) and fibroblast (30.0%) co-cultures. In a similar fashion, Wu et al. co-cultured pre-antral pig follicles with different follicular cells [86]. In contrast to Itoh and Hoshi’s results, Wu et al. found that small pre-antral follicle growth and survival was inhibited by co-culture with multiple follicles (with or without oocytes). The growth and survival of these follicles was only enhanced when co-cultured with cumulus cells from antral follicles >3 mm in diameter. Moreover, Ramesh et al. co-cultured buffalo pre-antral follicles with different somatic cells (cumulus, granulosa, mesenchymal and epithelial) [87]. Co-culture with cumulus, granulosa, and mesenchymal cells resulted in better development, growth rate, and survival than the control and epithelial cells. Maximum growth and survival was achieved via co-culture with cumulus cells, which supports Wu’s results. Therefore, these studies clearly demonstrate the utility and effect of co-culture. Similar co-culture experiments have also been conducted using a hybrid 2D/3D approach, in which stromal cells were cultured in 2D with primary follicles cultured in the state-of-the-art 3D biomaterial scaffolds [11, 12] (Figure 30.3a). Co-culture with ovarian stromal cells improved growth and survival of immature follicles [11]. Co-culture with mouse embryonic fibroblasts (MEFs), a commonly used feeder cell and stromal cell equivalent, also supported the survival and growth of early stage follicles within alginate beads and also resulted in the production of metaphase_II oocytes [12]. Finally, the media conditioned by MEFs showed similar effects, suggesting a unidirectional diffusible signal is responsible for the increased viability of early stage follicles. Nevertheless, stromal cells have not yet been incorporated into a 3D biomaterial environment, in which they may function in a more physiologic manner. Hence, the impact of co-culture could be significantly enhanced by developing fully 3D co-culture systems.
Recently, novel biomaterials and scaffold fabrication techniques have been applied to develop more biomimetic ovarian follicle culture systems that include stromal components in 3D. For example, Laronda et al. used decellularized bovine ovarian tissue slices as a substrate to culture and eventually transplant a population of murine primary ovarian cells that included granulosa, theca and stromal cells [88]. When transplanted, the recellularized decelluarized ovarian tissue initiated puberty in ovariectomized mice [88]. The decellularized extracellular matrix material is a tissue-specific protein “skeleton,” and likely retains important signaling factors, however the porosity is immutable, making it challenging to repopulate the scaffold with cells [89]. This challenge is particularly prominent for follicle culture, for which maintaining the 3D architecture of the follicle is so critical to survival and function. In order to be useful for ovarian follicle culture, decellularized tissues will likely need to be further processed into hydrogel materials that are able to support follicular architecture. Another emerging technique in tissue engineering that has been applied to ovarian follicle culture is 3D printing. Laronda and Rutz et al. fabricated microporous gelatin scaffolds using direct extrusion 3D printing and then seeded the scaffolds with mechanically isolated ovarian follicles [30]. Due to the mechanical isolation, some ovarian stromal cells remained attached to the follicle and were seeded into the scaffold. Stromal cells proliferated and migrated along scaffold struts and secreted the extracellular matrix protein laminin, which anchored follicles to the 3D-printed scaffold. The tortuous porosity of the 3D-printed scaffold maintained the 3D architecture of the follicle and supported follicular growth, function, and ovulation without the need for enzymatic removal of the biomaterial.
The integration of stromal cells into three-dimensional culture systems is possible, but will require some adaptations of the culture system. The challenge will be to develop a hydrogel matrix that promotes stromal cell survival within the constraints of follicle growth. Unfortunately, stromal cells cannot be added directly to the alginate follicle culture system without considerable modifications. Alginate does not support cell attachment. However, this can be accomplished by covalently linking alginate to full extracellular matrix proteins, such as collagen, fibronectin and laminin or small peptides sequences from these proteins. Without these attachment sites the cells will undergo apoptosis or cell death. Alternatively, other biomaterials such as collagen, Matrigel, decellularized extracellular matrix or polyethylene glycol (PEG) could be employed for co-culture. Unquestionably, the modifications needed for stromal cell culture will also influence follicle growth. Hence, it will be difficult to find overlapping culture conditions for both stromal cells and follicles. For example, the concentration of stromal cells must be fine-tuned to allow adequate paracrine signaling while not starving the follicle of oxygen or nutrients. Thus, developing a three-dimensional co-culture system will be challenging, but it is definitely an obtainable goal.
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