Background

Oocytes provide the overwhelming majority of cytoplasmic building blocks required by the embryo and must undergo two rounds of error-free chromosome segregation to ensure normal embryonic development (Chaigne et al., 2017; Greaney et al., 2017). The first of these divisions, meiosis I (MI), is notoriously error-prone, especially with advancing female age; indeed, errors arising during oocyte MI account for a staggering 80-90% of human aneuploidy (Greaney et al., 2017). MI is also the division in which symmetry-breaking occurs to enable the oocyte to expel half of its chromosomes into a very small daughter cell (known as the polar body, PB), whilst holding on to the bulk of the cytoplasm to support later embryonic development (Chaigne et al., 2017). It is well established that oocyte numbers, ovulation, and oocyte quality are the three oocyte-specific parameters that determine fertility and pregnancy success (Homer, 2020). While there are specific biomarkers and diagnostic tools to test oocyte numbers and ovulation, there is no definitive diagnostic test for oocyte quality. Although human oocytes would be the ideal research material for investigating oocyte quality, extensive restrictions and ethical concerns have made them almost completely inaccessible for experimentation. Hence, the molecular basis of oocyte quality remains poorly understood and will benefit from more in-depth investigation of key aspects such as meiotic maturation, fertilisation, and embryonic development. Interrogating the roles of genes that regulate these processes has been accomplished using surrogate mammalian models such as the mouse, which has become the most widely studied model.

Breakthrough advancements in understanding key molecular regulators of MI chromosome segregation and asymmetrical division, as well as the oocyte’s vulnerability to aging, have been dependent on the ability to perform detailed analyses of oocytes in vitro. The applications of this protocol are wide-ranging and allow for post-transcriptional gene silencing (for example, using RNAi and morpholinos), conventional biochemical analyses such as Western blotting, and in-depth visual interrogation of intracellular structures by immunocytochemistry and time-lapse microscopy. Our laboratory has used all these techniques during in vitro culture to uncover numerous unique facets of oocyte regulation, such as spindle assembly checkpoint (SAC) regulation of MI chromosome segregation and its decline with aging (Homer et al., 2005; Gui and Homer, 2012; Riris et al., 2014), a unique oocyte DNA damage response (Subramanian et al., 2020) and new genetic insight into DNA damage-based premature ovarian aging (Subramanian et al., 2021), novel aspects of regulation at the G2-M boundary (Homer et al., 2009; Gui and Homer, 2013), the contribution of late post-anaphase events to asymmetrical division (Wei et al., 2018 and 2020), and key metabolic nodes involved in oocyte aging and potential therapeutics targeting these nodes to reverse poor oocyte quality (Bertoldo et al., 2020; Iljas and Homer, 2020; Iljas et al., 2020).

There are a number of challenges involved in studying oocytes. One major challenge is that oocytes cannot be propagated in the lab like many immortal cell lines and must be isolated as primary cells each time they are required for experimentation. Another challenge is the very long duration of MI, which lasts 6-12 h depending on the mouse strain (versus minutes for mitosis), and requires lengthy experiments. Yet another challenge is the large volume of the oocyte (~40-times larger than that of a somatic cell), which makes the identification of small cellular structures markedly more difficult. It is therefore critically important to develop consistent and reproducible techniques for oocyte isolation and culture.

Oocyte culture methods were first developed as early as the 1930’s (Moricard and de Fonbrune, 1937). Later, experiments developed a simple isotonic solution capable of sustaining oocyte maturation (Donahue, 1968 and 1970), which allowed oocytes to be observed during MI for the first time. More sophisticated culture media were subsequently developed (Chatot et al., 1989), and now, oocyte isolation and culture protocols form the basis of all in vitro techniques used for studying oocyte biology. Here, we detail a method involving hormonal stimulation and collection of ovaries, followed by isolation and in vitro culture of oocytes.

In this protocol, pregnant mare serum gonadotrophin (PMSG) is used to stimulate antral follicle development, thereby increasing the yield of oocytes isolated from the ovary and enabling 30-100 oocytes to be obtained from each mouse depending on the strain and age (Wei et al., 2018 and 2020; Subramanian et al., 2020 and 2021). By minimising exposure to light and temperature fluctuations, this method reduces both the damage caused by in vitro culture and the resulting artefacts.

This protocol describes the collection of fully grown oocytes arrested in prophase of MI, characterised by the presence of an intact nucleus, which in oocytes is referred to as the germinal vesicle (GV). Prophase I-arrest can be experimentally maintained using chemicals such as 3-isobutyl-1- methylxanthine (IBMX), which sustains high cyclic adenosine monophosphate (cAMP) levels, thereby preventing activation of the major M phase-promoting kinase, cyclin-dependent kinase 1 (Cdk1) (Zhao et al., 2014). Conversely, resumption of meiotic maturation can easily be induced by washing out IBMX, thereby providing a reversible method for manipulating M-phase entry. This approach also allows for the timed introduction of small molecule inhibitors to disrupt regulators at any number of key timepoints during the cell cycle.

The applications of this protocol are extensive and have been critical for advancing our understanding of the regulation of MI.