Molecular mechanisms determining mammalian oocyte quality with the treatment of cancer therapy

Abstract

Cancer is a global public health issue and remains one of the leading causes of death in the United States (1). It is estimated in the US in 2022, about 935,000 new cases of cancer will be diagnosed in women, and the probability of developing invasive cancer is 5.8% for females younger than 50 years old (1). However, advances in screening programs, diagnostic methods, and therapeutic options have greatly increased the five-year survival rate in reproductive-age women with a variety of cancers. Given the clinical consequences of gonadotoxic cancer therapies, young, female cancer survivors may face compromised fertility, premature ovarian insufficiency, early-onset menopause, and endocrine dysregulation (2). Gonadotoxic side effects may include decreased oocyte quality within surviving follicles, loss of ovarian follicles, and impaired ovarian function. In reproductive-age women, oocyte quality is an important element for successful clinical pregnancies and healthy offspring as poor-quality oocytes may be a cause of infertility (3–5). Thus, it is critical to determine the quantity and quality of surviving follicles in the ovary after cancer treatment and to assess oocyte quality within those surviving follicles as these are markers for determining the capacity for ovarian function restoration and future fertility, especially for young cancer survivors (6). The long-term effects of cancer therapeutics on oocyte quality are influenced by factors including, but not limited to, individual patient characteristics (e.g. age, health history, comorbidities, etc.), disease type, or treatment regimen (7). These effects may translate clinically into an impaired production of viable oocytes and compromised fertility (8).

1. Cancer Therapy and Oocyte Quality

1.1. Radiation Therapy

Radiation therapy along with chemotherapy are cornerstones of cancer therapy and remain widely used as first or second line treatments of various malignancies (e.g., leukemia, Hodgkin’s disease, lymphomas, sarcomas, medulloblastomas, etc.) (7, 9, 10). However, during radiotherapy, healthy tissues near the tumor may be unavoidably exposed to radiations. In particular, radiation-induced damage to the ovaries is often irreversible and progressive (11). While radiotherapy destroys cancer cells by causing direct DNA damage as rapidly proliferating cancer cells with high mitotic activity and active DNA replication are more sensitive to radiation compared to cells with low mitotic division activity, oocytes appear to be an exception (7, 12). Human oocytes are generally extremely sensitive to radiation despite being arrested at prophase I of meiosis, and radiotherapy has been found to detrimentally impact ovarian function (7, 13). The extent of radiation-induced damage to the ovaries can vary depending on factors such as exposure dose, patient age, exposure time, and associated treatments.

Findings with female participants of the Childhood Cancer Survivor Study demonstrate a dose-dependent effect of radiation on the relative risk (RR) of ever being pregnant with the risk of pregnancy decreasing as the dose of ovarian/uterine radiation increases (5–10 Gray (Gy), RR=0.56, 95% confidence interval (CI)=0.37 to 0.85; >10 Gy, RR=0.18, 95% CI=0.13 to 0.26) (14). The study concluded that female cancer survivors (15 to 44 years old) exposed to an ovarian/uterine radiation dose greater than 5 Gy were less likely to become pregnant compared to sibling control participants (14). Ovarian exposure to radiation as low as 1 Gy was also found to be associated with greater risk for nonsurgical, premature menopause and premature acute ovarian failure (15, 16). One study with premenopausal cancer patients (<30 years old at the time of irradiation) reported that all patients who received radiation doses >15 Gy to the ovaries developed ovarian failure with increased serum levels of lactate dehydrogenase, prolactin, and follicle-stimulating hormone (FSH) and decreased serum levels of estradiol and progesterone, as markers of ovarian function (17). About half of the female patients who received pelvic irradiation that excluded at least one ovary using oophoropexy also developed ovarian dysfunction due to the additional effects of chemotherapy (17).

All women within the Georgia Cancer Registry who received pelvic radiotherapy (with or without chemotherapy) for a variety of malignancies experienced treatment-related amenorrhea (18). Furthermore, resumption of menses occurred later for women who were older at cancer diagnosis compared to women 20 to 24 years old (18). In a retrospective series with premenopausal (≤40 years old) cancer patients, treatment-related, long-term amenorrhea occurred in 4.2% of colon cancer patients and 94.1% of rectal cancer patients in part due to pelvic irradiation (19). Another study assessed ovarian function amongst a cohort of 100 female cancer survivors from the Childhood Cancer Registry who were treated with radiotherapy and/or chemotherapy (20). The authors found that at study onset, 17 survivors presented with premature ovarian failure and follicle-depleted or nondetectable ovaries along with elevated levels of FSH and luteinizing hormone (LH) (20). Furthermore, 70 survivors with preserved menstrual cycles had smaller ovarian volume (p<0.001) and a lower number of antral follicles per ovary (p<0.001) compared to controls along with increased estradiol levels (20). The total antral follicle count (AFC) was reduced with ovarian irradiation, exposure to alkylating chemotherapy, and older age at diagnosis (20). Taken together, these changes in hormone levels, ovarian volume, and follicle count compared to controls suggest impairment of ovarian potential despite the presence of preserved menses (20, 21).

Patients with primary or metastatic brain tumors who receive radiotherapy may also be at risk for infertility and ovarian dysfunction-related disorders as cranial irradiation can disrupt the hypothalamus-pituitary-gonadal axis (13). A questionnaire-based study with the German Childhood Cancer Registry evaluated fertility characteristics in childhood, brain tumor survivors who had been treated with cranial irradiation, specifically to the hypothalamic-pituitary-axis. The authors found that female participants who had received a radiation dosage of ≥30 Gy to the hypothalamic-pituitary-axis compared to dosages of 0–17 Gy and 18–29 Gy reported a higher frequency of permanent amenorrhea (p<0.001) and fewer pregnancies (p=0.008) (22). These findings are in accordance with earlier studies of female participants of the childhood cancer survivor study in the US where hypothalamic/pituitary irradiation doses of 22–27 Gy and > 30 Gy was associated with impaired fertility or was a significant risk factor for not having a pregnancy (23, 24). In one study, female patients treated with conformal radiation techniques for neoplasms arising from the base of skull had a 38% five-year risk of developing central hypogonadism in two to eleven years after radiation treatment (25). This indicates that the risk of endocrinopathy and infertility remains for female cancer patients even years after diagnosis and radiation treatment. Precocious puberty can also occur after low doses of cranial irradiation (18–24 Gy) in young girls being treated for brain tumors where future consequences may include hormone deficiencies and infertility (26). Furthermore, the relative risk of miscarriage is increased among women who received cranial or craniospinal irradiation compared to women without radiation treatment (27). Taken together, the findings from these studies suggest that even though follicles survive post-cancer treatment, the oocyte quality within surviving follicles may have dramatically decreased, resulting in lower pregnancy outcomes.

1.2. Chemotherapy

Radiotherapy is frequently used in combination with chemotherapy which can augment the risk of ovarian toxicity in cancer patients (13). Chemotherapy can be used alone or with other cancer therapeutics to treat an extensive group of diseases, and treatment includes a range of drug classes and combinations. Each class of chemotherapeutic agents has different mechanisms of action which ultimately halt the cell division cycle (2). There are many diverse treatment protocols that vary for different malignancies, and each has its individual risk of gonadotoxicity, with alkylating or alkylating-like agents deemed as high risk (28, 29). Furthermore, the extent of chemotherapy-induced gonadotoxicity depends on a range of factors like individual patient history, treatment protocol (e.g., dosage, drug class, drug combinations), or type of malignancy.

In a number of findings, the level of gonadotoxicity from chemotherapy appears to increase in a dose-dependent manner. For example, an earlier study found that childhood cancer survivors have an increased risk of developing nonsurgical, premature menopause compared to women who had never had cancer, and this risk increases with increasing dosage or exposure to radiation and/or alkylating chemotherapeutic drugs (16). Another report with a cohort from the Childhood Cancer Survivor Study demonstrated that exposure to increasing doses of alkylating agents was associated with decreased fertility and likelihood of becoming pregnant (14). A cross-sectional analysis of data from a prospective cohort study described significantly impaired reproductive hormone levels and decreased AFCs in cancer survivors compared to age-matched controls, supporting sub-clinical follicular depletion (30). Additionally, hormone levels in menstruating survivors who were exposed to high-dose therapy (alkylating agents, pelvic radiotherapy, or bone marrow transplant with total body irradiation) were similar to those of late-reproductive-age women (40 to 50 years old) (30). Other studies have found sustained chemotherapy-related amenorrhea and decreased AFCs in premenopausal cancer patients along with increased prevalence of menopause symptoms and significantly lowered spontaneous pregnancy rates in cancer patients treated high risk chemotherapy regimens (31, 32).

Older age has also been documented as an important risk factor for chemotherapy-induced ovarian damage using hormone levels and menstruation status as markers. One study cited a higher incidence of amenorrhea after chemotherapy treatment in breast cancer patients older than 40 years compared to younger patients (33). An earlier study stated that older age (34.7+8.0 years) at treatment initiation was associated with ovarian failure following chemotherapy as ovarian function was preserved in younger patients (27.4+8.3 years) after treatment (34). This may be due to a smaller follicle reserve in the ovaries of older women.

There have also been studies evaluating the efficacy of fertility therapies in patients previously exposed to cancer treatment. An early study found that fifteen women who underwent in vitro fertilization (IVF) after systemic cancer treatment (i.e. chemotherapy) had a poorer response to gonadotrophins compared to women who underwent localized cancer treatment but the number of retrieved oocytes was not statistically different (35). However, this study did not compare either group of cancer patients with a control group of women who did not have cancer (35). A later retrospective cohort study compared patients (<42 years) who had received chemotherapy and subsequently underwent fertility treatment with IVF or in vitro maturation (IVM) to age-matched patients (without cancer) undergoing the same fertility treatment protocol (36). The authors found that women previously treated for cancer with chemotherapy had a lower AFC and significantly decreased number of mature oocytes compared to the control group, demonstrating a reduced ovarian reserve as observed through decreased IVF/IVM outcomes (36). This is in accordance with another study demonstrating dramatically reduced IVF efficacy in cancer patients (22 to 33 years old) after undergoing one chemotherapy regimen; the number of retrieved oocytes was significantly lower (p<0.05) for cancer patients exposed to chemotherapy (average of 1.5 oocytes per patient) compared to the control group (average of 12.6 oocytes per patient) (37). Furthermore, only one good quality embryo was cryopreserved in the entire cancer-IVF patient group compared to the control group where four to eleven embryos per patient were obtained (37). As ovarian stimulation may involve oocytes that were exposed to chemotherapies during their growth phase, there are concerns regarding the genetic integrity and quality of those oocytes along with any embryos obtained from them.

More recent studies examined the live birth rates among women presenting for assisted reproductive technology. One study found that live birth rates among women using autologous oocytes decreased significantly for cancer survivors compared to women without cancer (47.7% without cancer versus 24.7% with cancer, p<0.0001), but there was no difference in live birth rates using donor oocytes among the two groups of women (38). These results indicate that factors acting in the pre- and peri-conceptional periods may have influenced the decline in live birth rates from autologous oocytes among cancer patients (38). In discordance with these findings, one study described similar live birth rates from autologous oocytes between cancer survivors and sibling controls within 22–48 weeks after a fresh IVF/ICSI (intracytoplasmic sperm injection) cycle (39). However, 47% of the IVF/ICSI treatments were for cancer survivors who were treated with surgery only, which is unlikely to damage the ovaries, leaving a moderately conserved ovarian reserve compared to survivors who underwent gonadotoxic cancer therapies (39). Outcomes such as decreased live birth rates, viable oocyte numbers, pregnancies, and infertility along with markers of ovarian impairment (e.g., amenorrhea, follicle counts, hormone levels) in patients exposed to chemotherapy can be taken as clinical manifestations of cancer therapy-induced ovarian damage and an indication of reduced oocyte quality.

1.3. Emerging Cancer Therapies

The gonadotoxic side effects of many conventional cancer treatments such as radiotherapy and chemotherapy have been well-documented in female cancer patients and survivors. However, the landscape for cancer therapeutics is rapidly changing, and there has been a transition to new treatment modalities such as targeted cancer therapy (40). Targeted therapy is more specific, more applicable to individualized care, and may be able to avoid off-target side effects compared to traditional cytotoxic cancer therapies (40). Many small-molecule inhibitors and immunotherapies, which are two classes of targeted therapy, are currently included in clinical oncological care. However, little is known regarding the potential ovotoxicity of these emerging cancer therapeutics and their long-term effects on reproduction for female cancer survivors (40).

Small-molecule inhibitors are low-molecular-weight compounds that can work at the cell surface or intracellularly and are designed to target specific oncogenic molecules implicated in tumor cell survival or cancer progression (40). Immunotherapy aims to alter the host immune system to enhance or restore the antitumor immune response. This can broadly be achieved by, but not limited to, encouraging pro-inflammatory states, increasing antigen presentation, and enhancing immune cell activation and specificity. In humans, studies characterizing the effects of small-molecule inhibitors and immunotherapy on ovarian function are generally limited to case reports of female menstruation, and amenorrhea is often reported as a primary outcome due to the ease of measurement (41).

Imatinib, pazopanib, lenvatinib, and inotuzumab ozogamicin are more novel therapeutic agents that have been reported to induce amenorrhea in premenopausal cancer patients (42–44). However, many of these patients were also exposed to different traditional chemotherapies. There have also been case reports of healthy conception and pregnancies in female patients who underwent treatment with small-molecule inhibitors and immunotherapies (45, 46). In general, there is a paucity of data and studies regarding the effect of targeted therapies on the ovaries and reproductive health of female cancer survivors. Thus, the effect of small-molecule inhibitors and immunotherapies on the quality of oocytes or embryos needs to be investigated.

2. Sensitivity to cancer therapy in ovarian reserve and oocyte quality

2.1. Formation of ovarian reserve and reproductive lifespan

Primordial follicle assembly is considered a critical aspect of female reproduction. Each primordial follicle consists of an oocyte at the prophase meiosis I and a single layer of flattened granulosa cells. The differentiation of germ cells into primary oocytes in the primordial follicle is poorly understood. In mice, at 10.5 days post coitus (dpc), primordial germ cells migrate from outside the embryo to the genital ridge and are then called oogonia once they colonize in the ovary. A cluster of up to 30 oogonia develops in a connected cluster and makes germ cell cysts. At 13.5 dpc, oogonia complete mitotic division and then become oocytes as they begin to enter meiosis.

At this time, oocytes transition through the prophase from leptotene to zygotene, to pachytene and arrest in the diplotene stage. The germ cells at diplotene arrest remain within a cyst, connecting with each other until beginning cyst breakdown. During cyst breakdown, the cluster of germ cells lose about two-thirds of their oocytes through apoptosis and the remaining oocytes are surrounded by pre-granulosa cells that form primordial follicles, consisting of ovarian reserve (47). The primordial follicle assembly process is coordinated by a variety of factors, including hormones, signaling molecules, and growth factors. As a result, appropriate coordination of primordial follicle assembly determines the quantity of ovarian reserve. Therefore, the regulation of primordial follicle assembly during the fetal and neonatal periods determines the long-term reproductive capacity of female mammals.

2.2. Primordial follicle damage

The DNA of oocytes, like other types of cells, is threatened by various forms of damage including radiation, chemotherapy, reactive oxygen species (ROS), and environmental toxicants (28). DNA damage may threaten genomic integrity in primordial follicles which remain quiescent in the ovary for decades. However, oocytes of primordial follicles are capable of executing an efficient repair program when faced with DNA damage. DNA double-strand breaks (DSBs) are considered to be the most toxic among DNA damages and are induced by irradiation and chemotherapy drugs such as cisplatin, cyclophosphamide, doxorubicin, etc. The repair mechanism of DNA DSBs in the oocytes has been well characterized (48), with two main mechanisms: non-homologous end jointing (NHEJ) (49) and homologous recombination (HR). NHEJ does not use a complementary DNA template and mediates the direct religation of broken DNA ends. There cannot be a relationship between NHEJ and the cell cycle, as NHEJ occurs mostly during G0/G1 phases. As a result, NHEJ is the most common repair mechanism for DNA damage in mitotic cells, but it is also error-prone (50). On the other hand, HR is the primary mode of DNA DSBs repair in meiotic cells because cells are mostly at the S and G2/M phases of the cell cycle, and sister chromatids are available as a template for accurate DNA repair (51). Both pathways can be functionally active in mammalian oocytes (52, 53). HR repair factors are located within oocytes of primordial follicles, suggesting that the HR repair mechanism is ready for DNA damage in immature oocytes. However, “decision-making” mechanisms involved in directing damaged cells toward HR or NHEJ are still under investigation. HR repair factors including RAD51, BRCA1, and BRCA2 are localized within oocytes of primordial follicles, suggesting that immature oocytes are equipped with the HR repair mechanism in case of DNA damage (54).

2.3. Damage from chemotherapy/radiation and underlying mechanism

As mentioned above, a great deal of our current understanding of the DNA damage response in primordial follicles has been developed as a result of studies exploring the effects of cancer therapy on fertility (55). It has been demonstrated that gonadotoxic therapies cause apoptotic death of primary oocytes via direct DNA damage by inducing DSBs (56). Survival of follicles is determined by the efficiency of DNA repair mechanisms. Severe DNA damage and inefficient repair result in primordial follicle death and decreased ovarian reserve.

At the molecular level, the elimination of primordial follicles following chemotherapy is mediated by the transactivation of the TAp63-related apoptotic pathway (57–60). p63, a member of the p53 family, plays an essential role in regulating maternal reproduction and genome integrity (61). Genes encoding p63 encode two major isoforms that are controlled by distinct promoters leading to transcripts differing in their N-terminus (62). Additionally, several variants are produced by alternative splicing at the C-terminus during transcription. Taken together, up to 10 diverse p63 isoforms exist in mammals, and usage of them is highly cell-type dependent. The longest one, TAp63α is highly expressed in the oocyte of the primordial follicle but disappears in the oocyte of the late secondary follicle (63).

Although p63 protein is highly expressed in oocytes of primordial follicles, cell death of primordial follicles may occur when p63 in their nuclei is phosphorylated by kinases that are activated by DNA damage, suggesting that p63 activity is tightly regulated (64). Accordingly, p63 remains in a close and dimeric conformation in oocytes. Through multistep phosphorylation, p63 is activated and triggers the apoptotic pathway. In response to DNA damage in particular, DNA DSBs kinases including Ataxia Telangiectasia mutated (ATM) (65) and Ataxia Telangiectasia and Rad3 related (ATR) are activated and phosphorylate several DNA damage response molecules such as a variant of the H2A protein family (H2AX) and checkpoint kinase 2 (CHK2) (56–58, 66, 67). In response to CHK2 phosphorylation, p63 is phosphorylated on S582, and casein kinase 1 (CK1) is recruited to add four phosphate groups sequentially. Therefore, DSBs trigger phosphorylation of p63, which leads to the formation of an open, active, and tetrameric conformation. Consequently, p63 in tetrameric conformation acts as a transcriptional activator in the transcription of two proapoptotic BH3-only proteins PUMA (BCL2 binding component 3; BBC3) and NOXA (phorbol-12-myristate-13-acetate induced protein 1; PMAIP1), which subsequently orchestrate the induction of apoptosis (68, 69).

The kinetics of p63 phosphorylation shows that it has the threshold for the induction of apoptosis (61). Therefore, to maintain genetic integrity, this activation mechanism must be adapted to a certain amount of damage that is low enough to protect the genetic pool, yet tolerant enough to not harm the reproductive system and initiate DNA damage repair system. Thus, if DNA damage does not surpass a certain threshold, the oocyte can be survived (61). One possible mechanism to explain the threshold of apoptosis via p63 is proposed that underphosphorylated p63 is degraded by proteasomal degradation. As a result of the fast proteasomal degradation of activated p63, this can be explained why the concentration of active isoforms of p63 is low within cells, whereas the concentration of inactive isoforms may be high. Consequently, when DNA damage is minor, fast degradation combined with slow activation could prevent apoptosis. Therefore, fast/slow kinetics ratios would determine the threshold for which DNA damage triggers apoptosis (61).

During evolution, this threshold level was established to prevent oocyte losses caused by accidental activation of p63 or low levels of DNA damage induced by reactive oxygen species in oocytes. In mice, ovarian reserves can eliminate with fewer than 10 DNA double-strand breaks per oocyte. In humans, the lethal dose for loss of the ovarian reserve was extrapolated to be less than 2Gy, while the typical total body irradiation dose for acute leukemia patients is 12Gy. In mice, a study has shown that 0.45 Gy of irradiation is sufficient to eliminate all oocytes expressing p63 through the ATM > CHK2 > TAp63α-hyper-phosphorylation pathway.

There are several ways in which chemotherapy can harm the ovary such as direct loss of primordial follicles, rapid activation of primordial follicles, follicular atresia, stromal damage, damage to the vasculature, or inflammation (28, 70, 71). However, different classes of chemotherapeutics exert different effects on the ovary. The type of DNA damage caused by each drug class may differ. For example, chemotherapeutics with cell cycle specificity have a primary effect only during specific phases of the cell cycle. Contrary to this, non-cell cycle-specific chemotherapeutics like alkylating agents (e.g., cyclophosphamide) can harm both proliferating and quiescent cells. As a result, these chemotherapeutic agents such as platinum-based agents and alkylating agents have the capacity to damage the DNA of diplotene-arrested oocytes.

2.4. Consequences of primordial follicle damage and clinical detection of ovarian reserve

Young cancer survivors often suffer from varying levels of gonadal impairment including primary ovarian insufficiency (POI) or diminished ovarian reserve, a weaker form of ovarian dysfunction. POI is defined as the loss of ovarian function before the age of 40 years which can cause infertility before the age of physiologic menopause. DOR is a condition characterized by retained regular menses and normal gonadotropins in the presence of biochemical markers.

The estimation of ovarian reserve is routinely performed through various ovarian reserve tests including FSH and anti-Mullerian hormone (AMH) and AFC (72). However, several studies have shown that AMH is more accurate at estimating ovarian reserve than AFC and baseline FSH. AMH is a serum marker for ovarian function that belongs to the transforming growth factor-beta family (73). Granulosa cells of the ovary express AMH from the primary follicle up to the small antral follicle. The number of these follicles indirectly reflects the number of remaining primordial follicles. Serum AMH levels fall rapidly following chemotherapy and re-detection of it in serum depends on the degree of ovarian damage. Thus, the AMH level indirectly reflects posttreatment ovarian function and oocyte quality as well. An accurate prediction of ovarian recovery can be made with the measurement of AMH at the end of chemotherapy. In this regard, undetectable AMH at the end of chemotherapy can be useful in identifying women who may not show ovarian recovery, which could influence the choice of endocrine therapy.

3. Communication between oocyte and granulosa cells to determine oocyte quality

3.1. Formation and physiological function of granulosa cells

In the ovary, the granulosa cell is the primary cell type that provides the physical support and microenvironment required for oocyte growth (74). The origin of granulosa cells has been resolved by tracking granulosa cells with the expression of forkhead transcription factor, FOXL2. Time-dependently knockout of foxl2 showed that the pre-granulosa cells are derived from the bipotential supporting cell precursors in the surface epithelium of mouse ovaries (75). A recent study using scRNAseq further revealed two distinct pathways of pre-granulosa cell differentiation. Bipotential pre-granulosa cells derived directly from bipotential precursors expressed Foxl2 early. A second pre-granulosa cell group, epithelial pre-granulosa cells, arise in the ovarian surface epithelium, ingresses cortically, and delays robust Foxl2 expression after birth. By embryonic day 19.5 (E19.5), epithelial pre-granulosa cells predominated in the cortex and differentiated into granulosa cells in primordial follicles. In contrast, medullar bipotential pre-granulosa cells differentiated along a distinct pathway to becoming wave 1 granulosa cells (76).

Oocyte growth is accompanied by granulosa cell growth and differentiation. Upon entry into the growth phase, the squamous pre-granulosa cells in primordial follicles become cuboidal and start mitotic proliferation to fully cover the expanding surface of the growing oocytes in primary follicles (77, 78). Granulosa cells continue to proliferate to generate multiple layers around the oocytes, and such follicles are termed secondary follicles. Theca cells are recruited to the external granulosa cell layers from Wt1-positive stromal cells and Gli1-positive mesonephros cells after birth in mice (79). As the oocytes grow and granulosa cells proliferate, a fluid-filled cavity termed the antrum appears, and such follicles are named antral follicles. At this stage, granulosa cells are differentiated into two subpopulations: the mural granulosa cells, which line the inner wall of the follicle, and the cumulus cells, which surround the oocytes. Under the opposing regulation of signals from intra- or extra-ovarian sources, the mural granulosa cells and cumulus cells express different genes and harbor different functions (80). Apart from anatomical differences in the location of the follicle, cumulus cells and mural granulosa cells are also functionally distinct (81). Cumulus cells support oocyte growth, produce hyaluronic acid, and undergo cumulus cell expansion in response to FSH. Mural granulosa cells usually carry out endocrine functions and support the development of the follicle (82). The size of follicles and phase of menstrual cycles affect the structure and function of cumulus cells and mural granulosa cells. In preovulatory follicles, with LH surge regulation, mural granulosa cells exhibit compact, dark cytoplasm and are shaped as epithelial-like cells. In contrast, cumulus cells show loose, clear cytoplasm with a round shape (83).

3.2. Granulosa cells and oocyte communication – importance to function as a follicle and to oocyte maturation

The communications between adjacent cells mainly take place via gap junctions, which allow the cell-to-cell molecular transfer (84). During follicle growth and development, gap junction-mediated communication between oocytes and granulosa cells is responsible for transporting nutritional molecules to oocytes and sharing metabolic intermediaries and products. Oocytes in growing follicles generate a proteinaceous matrix termed zona pellucida (ZP), which physically separates the granulosa cells from oocytes. ZP plays an important role in allowing only one sperm to penetrate the oocyte after insemination (85). Despite the separation from oocytes, granulosa cells maintain intimate connections with oocytes via transzonal projections (TZPs). TZPs are filopodia that are generated by granulosa cells regardless of the distance from oocytes and can bypass the proximal granulosa cells and transverse ZP (86, 87). Gap junctions are located at the feet of TZPs that come into contact with the oocyte.

Both gap junctions and TZPs offer the structural pathway for the exchange of molecules between granulosa cells and oocytes, and it was believed that the physiological direction of diffusive movement is from granulosa cells to oocytes. A recent study showed that the appropriate size markers injected into oocytes could diffuse into the granulosa cells through gap junctions and TZPs (87). Gap junctions and TZPs are responsible for trafficking metabolites and signaling molecules. Glycolysis and cholesterol metabolites are required for oocyte growth and development but cannot be produced in oocytes (88). Communication with granulosa cells is also necessary for oocytes to maintain a stable intracellular pH (89) and promotes chromatin remodeling and acquisition of meiotic competence (90, 91). The importance of gap junctions in oocyte development and maturation has been confirmed by mouse studies. Oocytes lacking connexin-37, a principal gap junction protein, failed to establish gap junctional communication with granulosa cells and exhibited impaired development, resulting in infertility (92, 93). Deleting connexin-43 in oocytes also abolished oocyte development, although oocyte-granulosa cell gap junctional communication remains intact (93–95). Besides gap junctions, granulosa cell-secreted KIT ligand has also been proposed to promote oocyte growth. The addition of soluble KIT ligand to culture medium increased the rate of oocyte growth without granulosa cells (96, 97). And blocking KIT by functional inhibitor suppressed oocyte growth in vitro (97). Deletion of Kit from growing oocytes would further explain the role of KIT during oocyte growth and maturation.

Paracrine growth factors secreted by oocytes stimulate metabolic pathways in granulosa cells, which are essential for oocyte growth and development, as well as granulosa cell proliferation and differentiation (81, 98, 99). The best-defined paracrine factors are members of the transforming growth factor-β (TGF-β) superfamily, including growth differentiation factors (GDFs) and bone morphogenetic proteins (BMPs), and fibroblast growth factor families (100–102). GDF9 is detectable in oocytes from primary follicles throughout the subsequent stages of folliculogenesis. Mice lacking Gdf9 were anovulatory and sterile with under-proliferated granulosa cells and decreased TZPs (103, 104). BMP15 is exclusively expressed in the oocytes but not critical for folliculogenesis (105). However, Bmp15 knockout female mice were subfertile with decreased ovulation and fertilization rates. This was exacerbated in Bmp15 ^−/−;Gdf9 ^+/− female mice, which were severely infertile due to abnormalities in ovarian folliculogenesis, cumulus cell physiology, and fertilization (106).

The intense cross-talk and co-operativity between the oocyte and the surrounding cumulus cells and mural cells is a major theme of oocyte maturation. Cumulus cells originate from a subpopulation of granulosa cells once the antrum is formed in the growing follicle (107). Oocyte–cumulus cell interactions are implemented through different signaling modalities and are recognized as a founding element of oocyte maturation. In fact, cumulus cells support a wide variety of functions of the maturing oocyte, including metabolism, meiotic arrest and resumption, and cytoskeletal rearrangements. Oocytes of primordial follicles are arrested at prophase I of meiosis until growth is completed and the preovulatory LH surge triggers meiotic maturation. This relies on cellular communication as LH receptors are not present in the oocyte but in mural granulosa cells and thecal cells. LH binds to its mural granulosa cell receptors and triggers the release of epidermal growth factor (EGF)-related peptides that diffuse through the antrum and bind to EGF receptors (EGFR) located on both the mural granulosa cells and cumulus cells (108). Cumulus cell expansion is triggered by the EGFR ligands that are released by the mural granulosa cells in response to LH. GDF9 and BMP15 also regulate cumulus cell expansion as the addition of GDF9 and/or BMP15 to culture medium induced cumulus expansion as well as the upregulation of their downstream(101, 102, 109) In addition, PTX3 has been studied to be essential in cumulus cell expansion in the process of oocyte maturation, ovulation, transportation through oviducts, and fertilization in mice and rats (110, 111).

3.3. Granulosa cells and Cancer Therapy

Besides the high sensitivity of primordial follicles to cancer treatment, granulosa cells are often impacted by chemo- or radiotherapies. Given the oocyte and its surrounding granulosa cells communicate bi-directly, cancer therapy-induced damage to granulosa cells will result in indirect damage to the oocyte, impacting oocyte quality and eventually affecting fertility. The impact on ovarian granulosa cells has been observed depending on different types of cancer treatments. TUNEL and caspase-3 positive ovarian granulosa cells were observed in doxorubicin (DOXO) injected mice, indicating the occurrence of apoptosis. A dramatic reduction in ovulation rate was also detected after DOXO injection and was not fully recovered after one month (112). A later study reported that double-strand DNA breaks following DOXO injection occurred in granulosa cells prior to DNA damage in oocytes (113). Consistently, DOXO caused DNA damage and apoptosis in both oocytes and granulosa cells of human follicles in an ovarian biopsy xenografted mouse model (114). Cyclophosphamide (CPA) triggered follicle loss by directly impacting oocytes in the ovarian reserve. Meanwhile, pyknosis and γH2AX signals were also observed in granulosa cells of growing follicles in mice after CPA exposure (115). Mitochondrial dysfunction and apoptosis in granulosa cells were induced in rats (116). CPA also caused significant damage in granulosa cell nuclei of human follicles after in vitro culture (117). Irinotecan, a topoisomerase I inhibitor, also induced apoptosis in granulosa cells after caspase-3 expression in growing follicles of mice (118).

In the clinic, a comparable density of viable antral follicles has been observed in patients exposed to chemotherapy, compared to non-exposed patients, especially at younger ages (119). However, the ovarian volume was significantly lower in patients who received gonadotoxic treatment (119). A deterioration in follicle quality was also reported after chemotherapy, with an increase in abnormal granulosa cell nuclei and oocyte vacuolization (120). As mentioned above, AMH is produced by the granulosa cells of growing follicles from the primary to the antral stage, and serum levels of AMH have been proposed to be used to reflect the ovarian reserve (121, 122). Serum AMH levels prior to cancer treatment have been investigated as a predictor of ovarian function after chemotherapy for breast cancer patients (123, 124). AMH levels aggressively dropped during chemotherapy in both prepubertal and pubertal girls, indicating the impairment of granulosa cells by chemotherapy (125). Radiotherapy also resulted in significantly lower AMH when targeted to pelvic/abdominal regions or total body irradiation (126–128).

3.4. Oocyte quality secondary to loss of function in granulosa cells

Granulosa cells are critical for ovarian function, including synthesizing steroid hormones and responding to hormonal signals, and cooperating with oocytes for follicle growth and maturation. Therefore, cancer therapy-induced damage to granulosa cells would impact oocyte quality. Current cancer therapies induce apoptosis in granulosa cells, which may have a negative effect on oocyte quality and pregnancy rates. It has been reported that the increased level of apoptosis in granulosa cells was associated with a reduction in oocyte quality, fertilization, pregnancy, and live birth rate in IVF patients (129–132).

Granulosa cells of growing follicles start to produce estradiol at the multilayer secondary follicles, and the estradiol level continues increasing with the proliferation of granulosa cells (133). Estradiol secreted from granulosa cells plays a positive feedback role in stimulating the production of FSH in the pituitary, which promotes further follicle growth and the production of steroid hormones in the ovary. Granulosa cells also produce activins, which are important in the development of early-stage follicles as autocrine and paracrine factors (134). Activins enhance FSH action on granulosa cells and inhibit androstenedione production by modulation of LH signals (135, 136). Luteinizing granulosa cells contain progesterone receptors that respond to the progesterone produced by corpus luteum cells, enhancing the activity of steroidogenic enzymes and response to FSH (137, 138). The damage of granulosa cells induced by cancer therapy dysregulates the secretion of endocrine hormones, eventually affecting oocyte quality.

4. Conclusion

Oocytes in the ovary can be directly affected by cancer therapy, causing loss of ovarian follicles and eventually infertility. If young cancer survivors maintain ovarian reserve post-treatment, oocytes may or may not carry DNA damage. In addition, oocytes can be indirectly targets affected by cancer therapy through granulosa or theca cells. Since oocytes are surrounded by granulosa and theca cells and communicate throughout oocyte maturation, damage to granulosa and theca cells can affect oocyte quality. Therefore, oocytes in young cancer survivors may not be comparable with ones from healthy young patients, as supported by lower pregnancy rates in cancer survivors. This demonstrates that oocyte quality in young cancer survivors is determined by multiple factors (Figure 1). The screening markers for oocyte quality are of utmost importance for patients who are exposed to cancer therapy.

Figure 1. Impact of cancer therapy on oocyte quality.

Ovarian follicles consist of oocytes and granulosa and theca cell layers during folliculogenesis in the ovary. Oocytes (Oo) communicate with granulosa cells (GCs) to develop into healthy follicles and are ready for producing hormones and ovulating oocytes. Thus, the communication between cells is key to developing high-quality oocytes for fertilization. However, cancer therapy can directly cause damage to oocytes and induce oocyte death via the TAp63-apoptotic pathway in primordial follicles (PF). In addition, it can indirectly cause somatic cell damage in GCs and theca cells and hinder the normal development of ovarian follicles because the communication between unhealthy cells (colored gray) is blocked or affects the growth of oocytes, especially growing follicles including antral follicles (AF). In this case, ovarian follicles cannot produce ovarian hormones, healthy oocytes, or high- quality oocytes, resulting in low pregnancy outcomes and infertility. Healthy versus unhealthy communication are indicated with arrows colored with purple and black, respectively. Healthy versus unhealthy granulosa cells are colored green and gray, respectively.

It has been shown and established clinically that conventional cancer treatments such as radiotherapy and chemotherapy are toxic to the ovary, and cancer survivors may have increased risks of premature menopause, early ovarian failure, endocrine disorders, and compromised fertility (2, 7, 28, 29, 41). With emerging targeted cancer therapies, the risk of reproductive toxicity in female cancer patients remains but is not as well characterized compared to traditional, cytotoxic treatment modalities (8, 40, 139, 140). Considering targeted cancer therapies are becoming more frequent in oncological care, further study is needed to evaluate class- and drug-specific effects on reproduction in female cancer patients and survivors. While various factors such as patient age, treatment regimen, or disease type influence reproductive health, the quantity and quality of the oocyte primarily determine fertilization success and the developmental fate of the embryo and fetus (3). Thus, cancer treatments that compromise both the quantity and quality of follicles along with oocytes will lead to ovarian toxicity (3). There are currently no effective pharmacological options to protect the ovaries of young girls and women receiving cancer treatment, but preserving oocyte quantity and quality before, during, or after cancer should be an important and fundamental goal.