Mammalian oocytes can be very long-lived cells and thereby are very likely to encounter DNA damage during their lifetime. Defective DNA repair may result in oocytes that are developmentally incompetent or give rise to progeny with congenital disorders. During oocyte maturation, damaged DNA is repaired primarily by non-homologous end joining (NHEJ) or homologous recombination (HR). Although these repair pathways have been studied extensively, the associated DNA synthesis is poorly characterized. Here, using porcine oocytes, we demonstrate that the DNA synthesis machinery is present during oocyte maturation and dynamically recruited to sites of DNA damage. DNA polymerase δ is identified as being crucial for oocyte DNA synthesis. Furthermore, inhibiting synthesis causes DNA damage to accumulate and delays the progression of oocyte maturation. Importantly, inhibition of the spindle assembly checkpoint (SAC) bypassed the delay of oocyte maturation caused by DNA synthesis inhibition. Finally, we found that ∼20% of unperturbed oocytes experienced spontaneously arising damage during maturation. Cumulatively, our findings indicate that oocyte maturation requires damage-associated DNA synthesis that is monitored by the SAC.

This article has an associated First Person interview with the first author of the paper.

Oocyte quality is a critical factor for female fertility and the transmission of high-quality genetic material to the offspring (Qiao et al., 2018). Defects in the oocyte genome can pose a serious threat to reproductive output (García-Rodríguez et al., 2018). Preserving DNA integrity during oocyte development is decisive and challenging (Katari et al., 2018). Mammals start life with a limited number of primary oocytes that progress through meiotic prophase I as far as the diplonema stage. At around birth, oocytes arrest in a quiescent phase called dictyate. Once the female reaches puberty, small clutches of these arrested oocytes will reactivate, progress from the germinal vesicle (GV) to metaphase II (MII) stage, and await fertilization before completing the second meiotic division (Mira, 1998). Oocytes are long-lived, and sensitive to chemo and radiotoxic stresses. Also, there is a high possibility of their being subject to genomic insults during their lifetime that may affect their development and maturation (Fragouli and Garrido, 2020). In addition, gamete aneuploidy is remarkably frequent in the female germline and is responsible for many pregnancies that end in miscarriages and births that carry a trisomy (Ménézo et al., 2010). Approximately 57% of pig oocytes, 30% of cattle oocytes and 30% of human oocytes show aneuploidy (Hornak et al., 2011). Therefore, oocyte quality is a critical factor in a successful fertility outcome.

During oocyte maturation, rapid detection and repair of the damaged DNA are critical for maintaining oocyte quality, avoiding genetic mutations, and ensuring accurate segregation at metaphase I (MI) and metaphase II (MII) (Mayer et al., 2016). The majority of DNA lesions (75%) are single-strand DNA (ssDNA) breaks, resulting from oxidative damage during metabolism or base hydrolysis. Furthermore, ssDNA breaks are converted into DNA double-strand breaks (DSBs), which are less frequent but more serious (Tubbs and Nussenzweig, 2017). Previous genomic and proteomic studies in mice and other vertebrates suggest the presence of base excision repair (BER), mismatch repair (MMR), nucleotide excision repair (NER), homologous recombination (HR) and non-homologous end-joining repair (NHEJ) machinery during oocyte maturation (Wang et al., 2010; Anastácio et al., 2017; Stringer et al., 2018).

The DNA damage response (DDR) in oocytes is similar to that of somatic cells, where checkpoint signaling is initiated by activation of the ataxia telangiectasia mutated (ATM) and ataxia telangiectasia and Rad3-related (ATR) protein kinases that phosphorylate downstream targets to amplify DDR signaling and either promote damage repair or initiate cell cycle arrest through spindle assembly checkpoint (SAC)-mediated inhibition of anaphase-promoting factor (Marangos et al., 2015; Macurek, 2016; Ma et al., 2019; Fragouli and Garrido, 2020; Ashwood-Smith and Edwards, 1996; Collins and Jones, 2016; Collins et al., 2015). Unlike what occurs in mice, extensive DNA damage does not induce cell cycle arrest in human oocytes. Instead, damaged oocytes can finish their maturation with abnormal spindles, and chromosome morphology indicates the differential regulation of oocyte DNA repair mechanisms in humans (Fragouli and Garrido, 2020; Rémillard-Labrosse et al., 2020). Previous results in mice have shown that exogenous induction of DNA damage activates HR and NHEJ (Martin et al., 2018; Qiao et al., 2018; Jin and Kim, 2017). HR is a multistep process where replication protein A (RPA) binds to 3′ ends of the ssDNA overhangs, created by nucleolytic resection, followed by RAD51 recruitment, resulting in the homology search and DNA strand invasion and exchange. The joint molecule formation is then followed by new DNA synthesis, which replaces nucleotides lost through DSB formation and 5′-terminal strand resection (Ashwood-Smith and Edwards, 1996; Hunter, 2015; Wright et al., 2018). In mouse oocytes, microinjection of recombinant RAD51, which plays a central role in HR, reduced the number of DNA breaks as well as the level of apoptosis (Kujjo et al., 2010; Bishop, 2012; Jin and Kim, 2017; Stringer et al., 2020; Bohrer et al., 2015). RAD51 is also necessary for doxorubicin-induced chemoresistance, and signifies active HR in fully grown mouse oocytes (Kujjo et al., 2010). Whereas NHEJ starts with recognizing and binding broken DNA ends by a heterodimer consisting of Ku70 and Ku80 (also known as XRCC6 and XRCC5, respectively). The DNA-dependent protein kinase catalytic subunit (DNA-PKcs; also known as PRKDC) is then recruited to Ku-bound broken DNA followed by nucleotide addition by the Pol X family polymerases. Finally, the DNA ligase IV complex ligates DNA breaks in both strands (Chang et al., 2017; Terasawa et al., 2014; Leem et al., 2019). Reports suggest that NHEJ components like ATM, Ku80 and DNA-PKcs are localized to DNA damage in mouse MII stage oocytes (Leem et al., 2019; Martin et al., 2018). Consistent with a role for NHEJ in DNA repair during oocyte maturation, pharmacological inhibition of either DNA-PKcs or ligase IV inhibits the DNA repair, indicating the presence of active NHEJ during the oocyte maturation (Martin et al., 2018). Although substantial steps in these DNA repair pathways are well studied, the nature and mechanisms of DNA synthesis during damaged DNA repair in maturing oocytes are, however, poorly characterized. Previously, Masui and Pedersen have shown that fully mature mouse oocytes exhibited unscheduled DNA synthesis when exposed to ultraviolet irradiation (Masui and Pedersen, 1975; Pedersen and Mangia, 1978). However, the molecular mechanisms are poorly understood. In this study, using porcine oocytes and chemical genetics, we have shown that the DNA synthesis machinery is readily available and recruited to the damage sites during maturation. The inhibition of the DNA synthesis in oocytes with chemically induced DNA damage limited the GV to MII progression due to the accumulation of the unrepaired DNA. Furthermore, knockdown of the SAC component Mad2 (also known as MAD2L1) suppressed maturation defects in damage-induced oocytes. Although chemically induced DNA damage was employed to study repair-associated DNA synthesis, we find that 20% of unperturbed oocytes experience spontaneous DNA damage. Thus, the damage-associated DNA synthesis repair system is necessary to maintain the oocyte quality and genome integrity.

Active DNA synthesis during oocyte maturation

To investigate the existence of DNA synthesis during the oocyte maturation, fully grown, GV staged, grade A, porcine oocytes were collected from antral follicles and cultured to MI and MII stages (Fig. S1) (Intawicha et al., 2019). GV, MI and MII stage oocytes were treated with 1 µg/ml etoposide to induce DNA damage and then assessed for DNA damage markers and de novo DNA synthesis via EdU incorporation (Fig. 1A–C). Bright EDU staining was observed and colocalized with γH2AX (the phosphorylated form of H2AX, a DNA damage maker; Hunter et al., 2001) staining in 70%, 81% and 96% of GV, MI and MII stage etoposide-treated oocytes, respectively. These results indicate that there is active DNA synthesis during damage-induced DNA repair and confirm the previous results of Masui and Pedersen, where they found the thymidine incorporation in UV-irradiated GV, MI and MII stage mouse oocytes (Fig. 1D) (Masui and Pedersen, 1975). Interestingly, ∼20% of untreated oocytes also showed EDU incorporation with γH2AX signals (Fig. 1D), suggesting that DNA synthesis may be active during oocyte maturation in spontaneously damaged oocytes (Pedersen and Mangia, 1978; Masui and Pedersen, 1975; Pedersen and Brandriff, 1980; Czołowska and Borsuk, 2000; Friedberg et al., 2014).

Fig. 1.

EdU incorporation in damage-induced oocytes. (A–C) Oocyte nuclei of respective stages immunostained for the DNA synthesis marker EdU (green), DNA damage marker γH2AX (red), and the DNA marker DAPI (magenta). (D) Quantification of the proportion (%) of oocytes where γH2AX signals colocalize with EdU staining in respective stages. GV, germinal vesicle-stage; MI, metaphase I; A/TI, anaphase/telophase I. Error bars show mean±s.e.m., n=40 to 60 nuclei for each stage. **P≤0.001; ***P≤0.0001 (unpaired two-tailed t-tests). Scale bars: 10 μm.

Fig. 1.

EdU incorporation in damage-induced oocytes. (A–C) Oocyte nuclei of respective stages immunostained for the DNA synthesis marker EdU (green), DNA damage marker γH2AX (red), and the DNA marker DAPI (magenta). (D) Quantification of the proportion (%) of oocytes where γH2AX signals colocalize with EdU staining in respective stages. GV, germinal vesicle-stage; MI, metaphase I; A/TI, anaphase/telophase I. Error bars show mean±s.e.m., n=40 to 60 nuclei for each stage. **P≤0.001; ***P≤0.0001 (unpaired two-tailed t-tests). Scale bars: 10 μm.

DNA synthesis is necessary for MI to MII progression in endogenously damaged and oocytes with exogenous damage induction

To test the requirement of DNA synthesis during oocyte DNA repair, GV stage oocytes were exposed to 1 µg/ml etoposide or 10 µM bleomycin, and then washed three times with fresh medium. After washing, oocytes were incubated with or without 50 mM DNA synthesis inhibitor hydroxyurea (HU) and cultured for 48 h (Fig. 2A,B; Fig. S3B). As previously reported, HU inhibits DNA synthesis by hindering the ribonucleotide reductase (Koç et al., 2004). The effect of inhibitors on oocyte maturation was evaluated by quantifying the proportion of cells with germinal vesicle breakdown (GVBD), and at the MI and MII stages based on their DNA morphology (Fig. S1). Approximately 70%, 75%, 52% and 70% oocytes matured to the MII stage with control, etoposide, HU and bleomycin treatments alone (Fig. S3B). Surprisingly, 95% of the HU with etoposide-treated and HU with bleomycin-treated oocytes were arrested before or at the MI stage (Fig. 2C; Fig. S3B). This inhibition is reversible, as the removal of the HU allowed oocytes to progress to MII (Fig. S4).

Fig. 2.

DNA damage following DNA synthesis inhibition causes MI arrest. (A) Schematic representation of oocyte treatments. (B) Oocyte nuclei of respective stages with respective treatment stained with the DNA marker DAPI (magenta) shown as a bright-field and fluorescence overlay. (C) Quantification of the proportion (%) of oocytes showing the DNA morphology indicative of the respective stages. GVBD, germinal vesicle breakdown; MI, metaphase I; A/TI, anaphase/telophase I; CTL, control; ETP, etoposide; HU, hydroxyurea; Aph, aphidicolin; ETP control, etoposide treatment control; ETP+HU, etoposide treatment with HU; ETP+APH, etoposide treatment with HU. Error bars show mean±s.e.m., n=40–60 oocytes for each treatment. *P≤0.01; n.s., P≥0.01 (unpaired two-tailed t-tests). Scale bars: 10 μm.

Fig. 2.

DNA damage following DNA synthesis inhibition causes MI arrest. (A) Schematic representation of oocyte treatments. (B) Oocyte nuclei of respective stages with respective treatment stained with the DNA marker DAPI (magenta) shown as a bright-field and fluorescence overlay. (C) Quantification of the proportion (%) of oocytes showing the DNA morphology indicative of the respective stages. GVBD, germinal vesicle breakdown; MI, metaphase I; A/TI, anaphase/telophase I; CTL, control; ETP, etoposide; HU, hydroxyurea; Aph, aphidicolin; ETP control, etoposide treatment control; ETP+HU, etoposide treatment with HU; ETP+APH, etoposide treatment with HU. Error bars show mean±s.e.m., n=40–60 oocytes for each treatment. *P≤0.01; n.s., P≥0.01 (unpaired two-tailed t-tests). Scale bars: 10 μm.

Furthermore, to eliminate the chance of HU-induced chemical toxicity in oocyte MI arrest, GV stage oocytes with DNA damage were exposed to 10 μM nucleoside analogue 2,3-dideoxycytidine (ddC) (Fig. S3A,B). As described previously, nucleoside analogs inhibit DNA synthesis by DNA chain termination (Dalakas et al., 2001). Consistent with the etoposide with HU inhibition oocyte maturation data, 80% of oocytes were arrested before or at the MI stage upon treatment with etoposide and ddC compared to 50% in ddC alone (Fig. S3B). These data imply that DNA synthesis is indispensable for the progression of MI to the MII stage but not the GV to the MI stage in DNA-damaged oocytes. Also, these results suggest that there is no robust G2 arrest in higher-order mammals (herein referring to mammals of higher-order than rodents) in response to DNA damage. Furthermore, to exclude the possibility of mitochondrial DNA synthesis effects, GV stage oocytes with chemically induced DNA damage were treated with or without 10 µg/ml aphidicolin (Fig. 2B). As previously reported, aphidicolin inhibits nuclear DNA polymerases but not the mitochondrial DNA polymerase, Pol γ (Baranovskiy et al., 2014). Aphidicolin treatment alone did not influence oocyte maturation. However, etoposide, followed by aphidicolin treatment, induced all the oocytes to arrest at GVBD and MI stages, indicating that nuclear DNA synthesis is crucial for DNA damage repair (Fig. 2C).

DNA synthesis is necessary for oocyte DNA repair

To further determine the reason for MI arrest in oocytes with chemically induced DNA damaged followed inhibited DNA synthesis oocytes, good quality GV stage oocytes were exposed to 1 µg/ml etoposide. They were then washed three times with fresh medium, followed by incubation with or without DNA synthesis inhibitors, cultured up to 48 h and monitored for DNA repair markers (Fig. 3A,B). γH2AX and RPA were present in control oocytes, and oocytes treated with etoposide or HU alone, only in the polar body nucleus, which degenerates through apoptosis, but not in the oocyte nucleus (Fabian et al., 2014; Wang et al., 2011) (Fig. 3B). However, RPA persisted in HU with etoposide- and HU with aphidicolin MI-arrested oocyte nucleus, and colocalized with γH2AX, indicating the accumulation of unrepaired DNA (Fig. 3B,C). These results suggest that DNA synthesis is necessary during oocyte DNA repair.

Fig. 3.

Accumulated breaks in DNA synthesis-inhibited oocytes. (A) Schematic representation of oocyte treatments. (B) Oocyte nuclei of corresponding treatments immunostained for DSB marker RPA (green) and the DNA damage marker γH2AX (red). Left panels are bright-field and fluorescence overlay; the two right-hand panels are fluorescence only. A dotted circle indicates the polar body. (C) Quantification of the proportion (%) of γH2AX- and RPA-positive nuclei for respective treatments. Error bars show mean±s.e.m., n=30 to 50 oocytes for each treatment. CTL, control; ETP, etoposide; HU, hydroxyurea; Aph, aphidicolin. **P≤0.001 (unpaired two-tailed t-tests). Scale bars: 10 μm.

Fig. 3.

Accumulated breaks in DNA synthesis-inhibited oocytes. (A) Schematic representation of oocyte treatments. (B) Oocyte nuclei of corresponding treatments immunostained for DSB marker RPA (green) and the DNA damage marker γH2AX (red). Left panels are bright-field and fluorescence overlay; the two right-hand panels are fluorescence only. A dotted circle indicates the polar body. (C) Quantification of the proportion (%) of γH2AX- and RPA-positive nuclei for respective treatments. Error bars show mean±s.e.m., n=30 to 50 oocytes for each treatment. CTL, control; ETP, etoposide; HU, hydroxyurea; Aph, aphidicolin. **P≤0.001 (unpaired two-tailed t-tests). Scale bars: 10 μm.

DNA synthesis machinery localizes to the repair sites during oocyte DNA repair

In eukaryotes, DNA replication starts with initiation, followed by DNA unwinding and polymerization (Duriez et al., 2019; Kelly and Callegari, 2019). Therefore, to investigate the DNA synthesis process during DNA repair, the MI stage damage-induced and non-induced oocytes were immunostained for origin licensing factor MCM2, which is known to interact with RAD51 to promote the ssDNA gap filling, clamp loader, PCNA, and its loading factor RFC1 (Fig. 4A–D) (Cabello-Lobato et al., 2021; Hamada et al., 2007; Kang et al., 2019). MCM2, PCNA and RFC1 are localized to the repair sites in 70% of damaged oocytes. Indeed, all the positively scored MCM2 and RFC1 foci colocalized with γH2AX (Fig. 4E). These results indicate that the DNA synthesis machinery is recruited to the damage sites during the oocyte DNA repair. Interestingly, ∼45% of control oocytes scored positively for PCNA. Out of them, 25% had PCNA signals that were not colocalized with γH2AX signals indicating the involvement of PCNA in other oocyte maturation processes (data not shown). Surprisingly, ∼15–20% of control oocytes were positive for MCM2 and RFC1 (Fig. 4E). Furthermore, to establish the connection between DNA repair and DNA synthesis, the MI stage oocytes that were subjected to chemically induced DNA damaged with or without aphidicolin were assessed for colocalization between the strand exchange protein RAD51 and PCNA loading replication factor RFC1 (Fig. 4F,G). Both RAD51 and RFC1 localized to nuclei in ∼60% of etoposide and aphidicolin co-treated oocytes compared to 20% in damage-induced oocytes (etoposide) or in oocytes treated with aphidicolin alone (Fig. 4H). Indeed most RAD51 patches in oocytes with RFC1, with a 0.82 positive correlation (Pearson correlation coefficient) at damage sites. These results indicate the recruitment of DNA synthesis machinery to the repair sites in DNA-damaged oocytes.

Fig. 4.

Recruitment of DNA synthesis machinery in damaged oocytes. (A,F) Schematic representation of oocyte treatments. (B–D) Etoposide-treated oocyte nuclei immunostained for γH2AX (red) and either PCNA or MCM2 or RFC1 (green) at the MI stage. (E) Quantification of the proportion (%) of PCNA-, MCM2- and RFC1-positive nuclei where PCNA, MCM2 and RFC1 colocalized with γH2AX signals in treated and control oocytes. (G) Control, etoposide-, aphidicolin- and etoposide with aphidicolin-treated oocyte nuclei immunostained for RAD51 (red) and RFC1 (green) at the MI stage. (H) Independent quantifications of the proportion (%) RAD51- and RFC1-positive nuclei in treated and control oocytes. Error bars show mean±s.e.m., n=30–50 oocytes for each treatment. *P≤0.01, **P≤0.001 (unpaired two-tailed t-tests). Scale bars: 10 μm.

Fig. 4.

Recruitment of DNA synthesis machinery in damaged oocytes. (A,F) Schematic representation of oocyte treatments. (B–D) Etoposide-treated oocyte nuclei immunostained for γH2AX (red) and either PCNA or MCM2 or RFC1 (green) at the MI stage. (E) Quantification of the proportion (%) of PCNA-, MCM2- and RFC1-positive nuclei where PCNA, MCM2 and RFC1 colocalized with γH2AX signals in treated and control oocytes. (G) Control, etoposide-, aphidicolin- and etoposide with aphidicolin-treated oocyte nuclei immunostained for RAD51 (red) and RFC1 (green) at the MI stage. (H) Independent quantifications of the proportion (%) RAD51- and RFC1-positive nuclei in treated and control oocytes. Error bars show mean±s.e.m., n=30–50 oocytes for each treatment. *P≤0.01, **P≤0.001 (unpaired two-tailed t-tests). Scale bars: 10 μm.

Dynamic localization of Pol β and δ during oocyte DNA repair

Mouse oocyte proteome studies of GV to MII stages have revealed the presence of Pol D1 and DNA synthesis machinery (Wang et al., 2010; Anastácio et al., 2017). Our data also indicates the requirement of DNA synthesis during oocyte maturation (Fig. 2B,C). Thus, to determine the polymerases involved in the DNA synthesis process, MI stage oocytes with and without chemically induced DNA damage were immunostained for DNA Pol β, δ, ε and γH2AX (Fig. 5A–D). Oocytes were assessed for the localization of the DNA polymerases and their colocalization with DNA damage marker γH2AX. DNA Pol β and δ localized to the chromatin and co-localized with γH2AX in 75% and 80% of the oocytes with DNA damage (Fig. 5E). However, ∼25% of control oocytes were positive for DNA Pol β and δ signals that colocalized with γH2AX (Fig. 5E). These results indicate the active participation of DNA Pol β and δ in damaged oocyte DNA repair. Surprisingly, DNA Pol ε localized to chromatin with or without DNA damage in all oocytes (Fig. 5D,E), indicating that Pol ε has other roles during oocyte maturation (see Discussion).

Fig. 5.

Recruitment of DNA polymerases in damaged oocytes. (A) Schematic representation of oocyte treatments. (B–D) Etoposide-treated oocyte nuclei immunostained for γH2AX (red) and either Pol β or δ or ε (green) at the MI stage. (E) Quantification of the proportion (%) of Pol β- or δ- or ε-positive nuclei where Pol β, δ or ε signals colocalized with γH2AX signals in treated and control oocytes. Error bars show mean±s.e.m., n=30–50 for control and etoposide treatment. *P≤0.01; n.s., P≥0.01 (unpaired two-tailed t-tests). Scale bars: 10 μm.

Fig. 5.

Recruitment of DNA polymerases in damaged oocytes. (A) Schematic representation of oocyte treatments. (B–D) Etoposide-treated oocyte nuclei immunostained for γH2AX (red) and either Pol β or δ or ε (green) at the MI stage. (E) Quantification of the proportion (%) of Pol β- or δ- or ε-positive nuclei where Pol β, δ or ε signals colocalized with γH2AX signals in treated and control oocytes. Error bars show mean±s.e.m., n=30–50 for control and etoposide treatment. *P≤0.01; n.s., P≥0.01 (unpaired two-tailed t-tests). Scale bars: 10 μm.

Pol δ is essential for oocyte DNA repair

To determine whether Pol δ and ε were required for oocyte maturation following DNA damage, oocytes with and without DNA damage were injected with negative control, PCNA, Pol δ and Pol ε siRNA (Fig. 6A,B). The effect of siRNA injection on oocyte maturation was assessed by quantifying the proportion of oocytes that reached the GVBD, MI and anaphase/telophase I (A/TI) stage by assessing the DNA structure. Except for the negative control, <20% of the PCNA and Pol δ siRNA-injected oocytes reached the A/TI stage (Fig. 6C), whereas 40% of Pol ε siRNA-injected oocytes reached the A/TI stage. These results indicate that PCNA and DNA Pol δ are critical during damaged oocyte DNA repair and oocyte maturation. In mouse oocytes, exogenous DNA damage induces MI arrest by normal spindles with a higher percentage of chromosome mislocalization or fragmentation (Collins et al., 2015; Marangos et al., 2015). Therefore, to understand the reason behind PCNA and Pol δ knockdown-induced MI arrest, we immunostained negative control, PCNA, Pol δ, and Pol ε siRNA-injected MI stage oocytes to assess spindle assembly and chromosome organization (Fig. 6D). Oocytes were assessed for the localization of the misaligned chromosomes away from the spindle. Approximately 50% of PCNA-, Pol δ- and Pol ε siRNA-injected oocytes had chromosome misalignment compared to 17% of the negative control (Fig. 6E).

Fig. 6.

MI arrest in oocytes with knockdown of DNA polymerases. (A) Schematic representation of oocyte treatments and microinjections. (B) Negative (-ve) control or Pol δ siRNA microinjected oocyte nuclei stained for the DNA marker DAPI (magenta) shown as a bright-field and fluorescence overlay. (C) Quantification of the proportion (%) of oocytes at A/TI as assessed by DNA morphology after respective siRNA microinjections. (D) siRNA microinjected oocyte nuclei stained for spindle (green) and chromatin (magenta) shown as a bright-field and fluorescence overlay. Arrows indicate the misaligned chromosomes. (E) Quantification of proportion (%) of oocytes with fragmented chromatin in negative control and respective siRNA microinjected oocytes. A/TI, anaphase/telophase I. Error bars show mean±s.e.m., n=50–70 oocytes injected with respective siRNA. **P≤0.001, *P≤0.01 (unpaired two-tailed t-tests). Scale bars: 10 μm.

Fig. 6.

MI arrest in oocytes with knockdown of DNA polymerases. (A) Schematic representation of oocyte treatments and microinjections. (B) Negative (-ve) control or Pol δ siRNA microinjected oocyte nuclei stained for the DNA marker DAPI (magenta) shown as a bright-field and fluorescence overlay. (C) Quantification of the proportion (%) of oocytes at A/TI as assessed by DNA morphology after respective siRNA microinjections. (D) siRNA microinjected oocyte nuclei stained for spindle (green) and chromatin (magenta) shown as a bright-field and fluorescence overlay. Arrows indicate the misaligned chromosomes. (E) Quantification of proportion (%) of oocytes with fragmented chromatin in negative control and respective siRNA microinjected oocytes. A/TI, anaphase/telophase I. Error bars show mean±s.e.m., n=50–70 oocytes injected with respective siRNA. **P≤0.001, *P≤0.01 (unpaired two-tailed t-tests). Scale bars: 10 μm.

Defects in DNA synthesis induces SAC in damaged oocytes

Previous reports suggest that SAC-mediated DNA-damage checkpoint response induces MI arrest in mouse oocytes (Marangos et al., 2015; Collins et al., 2015). Therefore, to test SAC-mediated checkpoint mechanisms in higher-order mammals, MI-arrested oocytes were immunostained for the SAC marker Mad2 (Fig. 7A,B). As previously reported, Mad2 is the major component, along with Bub1, BubR1 and Mps1, that accumulates on unattached kinetochores to inhibit the APC/C (Marangos et al., 2015; Collins et al., 2015). Mad2 persisted on chromatin in ∼60% of oocytes with DNA damage followed by DNA synthesis inhibition, compared to controls (Fig. 7C). Furthermore, to determine whether the MI arrest is really because of accumulated Mad2, oocytes with DNA damage were injected with negative control, Pol δ or Mad2 siRNA alone, and with Pol δ with Mad2 siRNAs together (Fig. 7D,E). Approximately 30% more Pol δ siRNA-injected oocytes were arrested at MI than those with negative control and Mad2 siRNA. However, the loss of Mad2 in the Pol δ siRNA background rescues the maturation defects in Pol δ siRNA-injected oocytes (Fig. 7F). These results imply that the active SAC controls the DNA damage-induced MI arrest in higher-order mammals.

Fig. 7.

SAC-dependent damage-induced MI arrest. (A,D) Schematic representation of oocyte treatments and microinjections. (B) Etoposide- and etoposide with HU-treated oocyte nuclei stained for Mad2 (green) and DNA (magenta) at the MI stage shown as a bright-field and fluorescence overlay. (C) Quantification of the proportion (%) of Mad2-positive nuclei in etoposide- and etoposide with HU-treated oocytes. (E) Oocyte nuclei of respective stages with respective microinjections stained for DNA (magenta) shown as a bright-field and fluorescence overlay. (F) Quantification of the proportion (%) of oocytes at A/TI as assessed by DNA morphology after respective treatment with etoposide, and etoposide with HU. A/TI, anaphase/telophase I; ETP, etoposide; HU, hydroxyurea. Error bars show mean±s.e.m., n=40 oocytes were analyzed for Mad2 localization for each treatment (B); n=30–50 oocytes were injected with respective siRNA (F). **P≤0.001; *P≤0.01; n.s., P≥0.01 (unpaired two-tailed t-tests). Scale bars: 10 μm.

Fig. 7.

SAC-dependent damage-induced MI arrest. (A,D) Schematic representation of oocyte treatments and microinjections. (B) Etoposide- and etoposide with HU-treated oocyte nuclei stained for Mad2 (green) and DNA (magenta) at the MI stage shown as a bright-field and fluorescence overlay. (C) Quantification of the proportion (%) of Mad2-positive nuclei in etoposide- and etoposide with HU-treated oocytes. (E) Oocyte nuclei of respective stages with respective microinjections stained for DNA (magenta) shown as a bright-field and fluorescence overlay. (F) Quantification of the proportion (%) of oocytes at A/TI as assessed by DNA morphology after respective treatment with etoposide, and etoposide with HU. A/TI, anaphase/telophase I; ETP, etoposide; HU, hydroxyurea. Error bars show mean±s.e.m., n=40 oocytes were analyzed for Mad2 localization for each treatment (B); n=30–50 oocytes were injected with respective siRNA (F). **P≤0.001; *P≤0.01; n.s., P≥0.01 (unpaired two-tailed t-tests). Scale bars: 10 μm.

Spontaneously induced endogenous cellular processes reduce oocyte quality

The increased ROS levels during the ovarian aging affects the oocyte quality by inducing DNA damage (Homer, 2021). Therefore, to test whether increased ROS contribute to the 20% of unperturbed oocytes that showed spontaneous DNA damage, we analyzed the ROS, γH2AX, and RFC1 levels in freshly collected GV stage oocytes (Fig. 8A,B). We scored three classes of ROS based on the signal intensities (comparative intensity mean value of oocyte area): (1) oocytes with low ROS having signal intensities 0.1 to 0.5; (2) oocytes with medium ROS having signal intensities above 0.5 to 1; and (3) oocytes with very high ROS having signal intensities more than 1 (Fig. 8A,B). The majority of the freshly isolated oocytes scored for class 1. Surprisingly, 17% of oocytes scored for class 2, and another 20% score for class 3, indicating that the spontaneously induced endogenous cellular processes may cause the high levels of ROS seen in 20% of the oocytes (Fig. 8C). Like ROS signals, the majority of freshly isolated oocytes were negative for RFC1 and γH2AX. However, most ROS oocytes with high ROS level also had γH2AX and RFC1 signals, with a positive correlation of 0.72 and 0.81 (Pearson's correlation coefficient), indicating spontaneously induced endogenous cellular processes, like oxidative stresses, may contribute to oocyte DNA damage (Fig. 8A,D). Also, the positive correlation of RFC1 with ROS indicates local DNA synthesis mechanisms during oocyte repair (Fig. 8B,D).

Fig. 8.

ROS levels in freshly isolated oocytes. (A,B) Freshly isolated oocytes immunostained for ROS (green), the DNA damage marker γH2AX or PCNA loading factor RFC1 (red), and DNA (magenta). The dotted circles labeled with 1 indicates the GV stage oocyte with >1 ROS signal intensity and colocalization with γH2AX or RFC1. The dotted circles labeled with 2 indicates the GV stage oocyte with 0.1–0.5 ROS signal intensity and colocalization with γH2AX or RFC1. (C) Quantification of the proportion (%) of ROS signal intensities seen in freshly isolated GV stage oocytes. (D) Fraction of different ROS classes (1, green; 2, orange; 3, red; see text) oocytes that were positive for γH2AX or RFC1. Error bars shows mean±s.e.m., n=60–100 oocytes were analyzed for ROS in each animal. Scale bars: 10 μm.

Fig. 8.

ROS levels in freshly isolated oocytes. (A,B) Freshly isolated oocytes immunostained for ROS (green), the DNA damage marker γH2AX or PCNA loading factor RFC1 (red), and DNA (magenta). The dotted circles labeled with 1 indicates the GV stage oocyte with >1 ROS signal intensity and colocalization with γH2AX or RFC1. The dotted circles labeled with 2 indicates the GV stage oocyte with 0.1–0.5 ROS signal intensity and colocalization with γH2AX or RFC1. (C) Quantification of the proportion (%) of ROS signal intensities seen in freshly isolated GV stage oocytes. (D) Fraction of different ROS classes (1, green; 2, orange; 3, red; see text) oocytes that were positive for γH2AX or RFC1. Error bars shows mean±s.e.m., n=60–100 oocytes were analyzed for ROS in each animal. Scale bars: 10 μm.

Oocyte quality is a critical factor for female fertility and transmission of high-quality genetic material to the offspring (Wang and Höög, 2006; Stringer et al., 2020, 2018; Martin et al., 2019). Hence, it is important to maintain the quality of the oocyte through responses to different stresses, which causes DNA damage. Like somatic cells, oocytes have specific mechanisms to detect and respond to the genomic insults they come across. Several studies have concluded that the oocytes exposed to exogenous chemical agents activate the HR and NHEJ repair pathways (Leem et al., 2019; Martin et al., 2018; Ma et al., 2019; Kujjo et al., 2012, 2010; Jin and Kim, 2017; Gruhn et al., 2013). However, the roles of DNA synthesis during repair remain unanswered. Previous studies have shown that aneuploidy in higher-order mammals occurs more frequently than in the mouse (Hornak et al., 2011; Rémillard-Labrosse et al., 2020). Thus, in this study using porcine oocytes, we have demonstrated that DNA synthesis mechanisms are critical for oocyte maturation and genome stability.

The necessity of nuclear DNA damage-associated DNA synthesis during oocyte maturation

Previous studies in mice have indicated that DNA synthesis is active when the oocytes are exposed to ultraviolet radiation (Pedersen and Mangia, 1978; Masui and Pedersen, 1975; Czołowska and Borsuk, 2000). Moreover, the systematic proteome profiling of mouse oocytes at different stages during the maturation has revealed the presence of DNA repair and synthesis machinery (Wang et al., 2010; Anastácio et al., 2017). In our study, to understand the importance of DNA synthesis during DNA repair, a lower concentration of etoposide treatment was used to generate minor DNA damage compared to previous studies in porcine oocytes (Wang et al., 2016). We found that 1 µg/ml etoposide for 1 h treatment enough to generate the minimum number of DNA breaks to initiate DNA synthesis, without compromising the meiotic progression (Fig. S2). Oocytes with DNA damage that were then subjected to an inhibition of DNA synthesis, arrested at MI with unrepaired DNA, indicating that DNA synthesis is required during oocyte repair. Furthermore, we found that the recruitment of the PCNA, MCM2, RFC1, and Pol δ to the repair sites, confirming that DNA synthesis occurs during oocyte repair. Then the question arises of why and how is the DNA synthesized during the repair. For instance, if an oocyte encounters DNA damage in the form of DSBs in GV, MI or MII stages, is the DDR in oocytes is initiated by activating the ATM and ATR kinases? Furthermore, ATM and ATR phosphorylates the downstream targets Chk1 and Chk2 to amplify DDR signaling, allowing the oocytes to choose the DNA repair pathways. In oocytes, where the HR pathway is extensively studied, the generation of ssDNA follows the recruitment of RPA, and Rad51 facilitates the strand invasion to form a joint molecule. Finally, the gap is filled by synthesizing the new piece of DNA (Ma et al., 2021). Previous studies in Xenopus oocytes, where UV-damaged plasmid DNA was shown to be repaired by excision repair through DNA Pol δ raises the possibility of active participation of other DNA repair processes like NER, BER, MMR and NHEJ during porcine oocyte DNA repair (Sweigert and Carroll, 1990; Ackerman et al., 1999; Varlet et al., 1996; Lin et al., 2019; Yagi et al., 2005; Matsumoto, 1999). Detailed experiments are necessary to study individual pathways and their requirements. In contrast to DNA Pol δ, the Pol ε localization in damaged and non-damaged oocytes is independent of DNA damage. In plants, DNA Pol ε plays a significant role in heterochromatin organization and maintenance (Li and Zhang, 2012). Further studies are necessary to delineate the functions of Pol ε during oocyte maturation.

Weak G2/M checkpoint in higher-order mammalian oocytes

In our current study, we exposed the oocytes to the DNA-damaging agents at the GV stage and then introduced DNA synthesis inhibitors to evaluate the requirements of DNA synthesis during maturation. Most of the etoposide with HU- and bleomycin with HU-treated oocytes were arrested at the MI stage without a G2/M arrest. The G2/M checkpoint is a vital barrier that restricts the M phase entry of DNA-damaged cells until the damage is repaired (Subramanian et al., 2020). Unlike what is seen in mice, higher-order mammalian oocytes spend most of their time in GV to MI transition (Fig. S1). For example, the GV to MI transition in mice is 2 to 4 h, in pig, it is 24 to 30 h, and in humans it is 16 to 20 h (Conti and Franciosi, 2018; Miyano et al., 2003; Combelles et al., 2002; Sirard et al., 1989). However, there is no arrest point before or after GVBD. DDR in oocytes is initiated by autophosphorylation and activation of ATM/ATR kinases (see above). However, in porcine oocytes at the G2 stage, 25 µg/ml of etoposide induces high levels of DNA breaks but does not activate ATM/ATR kinase (Wang et al., 2016). In contrast, treatment with 50 and 100 µg/ml etoposide activates the ATM/ATR kinase but not Chk1, which could be one of the possible reasons for damaged oocytes arrest at MI but not at GV or GVBD (Marangos and Carroll, 2012; Rémillard-Labrosse et al., 2020). Surprisingly, ∼45% of etoposide with aphidicolin-treated oocytes arrest at GVBD-like stages, raising the possibility of a synergistic chemical effect that may cause a high level of DNA damage, which completely halts the cells at a GVBD-like stage.

How do oocytes respond to DNA insults during natural maturation in ovarian follicles?

DNA repair processes in oocytes have been extensively studied by exogenously exposing the oocytes to damage agents in culture medium (Marangos et al., 2015; Collins et al., 2015). However, how the oocytes respond to the DNA damage during natural maturation in ovarian follicles is unknown. The current study found that, naturally, ∼20% of freshly isolated GV-stage oocytes were positive for EdU staining that co-localized with the DNA damage marker γH2AX (Fig. 1D). We also found that the DNA repair protein RPA and DNA synthesis machinery RFC1 were present in 20 to 25% of fresh GV stage oocytes (Figs 3B and 4B–D). Consistent with our results, DNA repair and synthesis factors are present in the mouse GV stage oocyte proteome (Wang et al., 2010). These results indicate, in a natural setting in the ovarian follicles, damaged DNA could be repaired, like in in vitro maturation (IVM) conditions. Furthermore, we have shown that spontaneously induced endogenous cellular processes (oxidative stresses) may generate high levels of ROS, subsequently leading to DNA damage (Fig. 8A–C).

SAC-dependent and -independent MI arrest in DNA-damaged oocytes in higher-order mammals

In mouse oocytes, DNA damage induces SAC-mediated MI arrest (Marangos et al., 2015; Collins et al., 2015). However, human oocytes with DNA damage can finish MII, suggesting the absence of SAC-mediated arrest during oocyte maturation (Rémillard-Labrosse et al., 2020). These results raise the possibility of differential regulation of SAC-mediated MI arrest in higher-order vertebrate oocytes. In our study, we have found that the inhibition of MAD2 rescued the porcine oocytes from MI arrest. However, only 50% of the oocytes could complete MI, indicating that the remaining oocytes arrest independently of the SAC. Also, we cannot rule out the possibility of incomplete inhibition of MAD2 by siRNA or that cells failing to progress might undergo apoptosis. Further studies are necessary to separate SAC-dependent and -independent modes of damaged oocyte arrest at MI.

Spontaneously induced DNA assaults and their impact on oocyte quality and genome stability

Our present study raises the possibility that continuous endogenous stress in oocytes leads to DNA damage, which may give rise to mutations in the mitochondrial and nuclear genome leading to genome instability. We also found that the ROS levels increased in the oocytes with age (data not shown). ROS is one of the most significant endogenous stresses that cells come across. ROS and their byproducts produced from the aerobic metabolism causes mitochondrial and nuclear DNA damage (Kaludercic et al., 2014; Pizzino et al., 2017; Sasaki et al., 2019). ROS levels increase during the in vitro maturation of oocytes in all organisms (Mihalas et al., 2017). The addition of anti-oxidants subsequently increases the oocyte maturation efficiency and quality (Mihalas et al., 2017; Leem et al., 2019; Liang et al., 2017). Moreover, in vitro matured oocytes are more susceptible to exogenous damaging agents than in vivo matured oocytes (Uppangala et al., 2015). One of the critical factors in assisted reproductive technology is to get a large number of competent oocytes for the maturation and subsequent downstream processes (Swain and Pool, 2008). Therefore, anti-ROS agents might be beneficial for women undergoing assisted reproductive technology (ART) or pregnancy cycles, and further studies should look into this possibility.

Oocytes collection and culture

Porcine ovaries were collected from a nearby local slaughterhouse and transported to the laboratory within 30 min in a 50 ml tube containing PBS with 100 IU/ml penicillin and 0.05 mg/ml streptomycin. All animal experiments were performed according to Institutional Animal Ethics Committee (IAEC) guidelines. Oocytes were aspirated after puncturing and slicing the ovary in oocyte collection medium [TCM199 containing, 5.0 g/l BSA, and 50 µg/ml penicillin-streptomycin (Sigma, P4333)]. Only oocytes having a uniform ooplasm and compact three to four layers of cumulus cells were chosen for further experiments. After being washed three to four times in the same medium and once in the in vitro maturation (IVM) medium, which contains TCM199, 10 IU/ml follicle-stimulating hormone (FSH), 10 IU/ml luteinizing hormone (LH), 10% follicular fluid, 10% fetal bovine serum (FBS), and 50 µg/ml penicillin-streptomycin. Twenty oocytes were placed in 100 µl droplets of IVM medium, overlaid with mineral oil, and were cultured for 48 h at 37°C under 5% CO2 in humidified air. After maturation, the cumulus surrounding oocytes were denuded by pipetting in TCM199. The denuded oocytes were examined under a zoom stereo microscope (SMZ745; Nikon) for maturation. Oocytes showing a polar body were considered mature.

ROS measurement

To quantify the ROS levels, freshly isolated oocytes were incubated in IVM medium containing 10 μM 2′,7′ dichlorodihydrofluorescein diacetate (H2DCFDA; Invitrogen, D399) for 5 min, followed by three washes with PBS. They were examined under the fluorescence microscope (Axio Observer 7 - Carl Zeiss Microscopy GmbH) for Alexa Fluor (AF) 488. The quantification of fluorescence intensity was measured using Zeiss 2.6 blue edition software.

EdU labeling and staining

Nascent DNA synthesis was labeled and stained with Click-iT EdU Alexa Fluor 488 Imaging Kit (PK-CA724-488FM, PromoCell GmbH, Germany) according to the manufacturer's instructions with minor modifications. 20 µM EdU was added to the IVM medium for at least 6 h at three different time points (i.e. 0 h, 30 h and 48 h). After labeling, oocytes were washed three times with PBS followed by fixation with 4% paraformaldehyde (PFA) and 0.05% Triton X-100 (Sigma, T8787) on a glass slide. After fixation, oocytes were rinsed three times with Tris-buffered saline with 0.05% Triton X-100 (TBST) and then stained with anti-γH2AX primary antibody (Millipore, 05-363-I, 1:500), dilution 1:500, overnight at room temperature in a humid chamber. The next day, the oocytes were washed three times with TBST, blocked with blocking buffer for 30 min, and incubated with AF 555-conjugated secondary antibodies, diluted at 1:2000 in antibody dilution buffer (ADB) for 2 h at room temperature in a humidified chamber. Then the oocytes were washed twice with TBST after the EdU reaction was carried out in the dark for 30 min as per the instructions. Finally, oocytes were counterstained with DAPI, mounted with Prolong Diamond, and imaged by fluorescence microscopy.

Chemical treatments

To induce the minor DNA damage, freshly retrieved GV stage oocytes were treated with 1, 5 and 10 µg/ml of etoposide (Sigma, A1383). Based on the results, 1 µg/ml was selected for the rest of the studies. Furthermore, the oocytes were exposed to the following chemical treatments. First, etoposide followed by HU treatment. For this, freshly retrieved GV stage oocytes were cultured in IVM medium supplemented with 1 µg/ml etoposide for 1 h in 5% CO2 with humidified air, followed by three washes with TCM199, then culture for 48 h in IVM medium with and without 50 mM HU. Second, bleomycin followed by HU treatment. For this, freshly retrieved GV stage oocytes were cultured in IVM medium supplemented with 10 µM bleomycin (SRL, 20899) for 1 h in 5% CO2 with humidified air, followed by three washes with TCM199, then cultured for 48 h in IVM medium with and without 50 mM HU. Three, etoposide followed by aphidicolin treatment. For this, freshly retrieved GV stage oocytes were cultured in IVM medium supplemented with 1 µg/ml etoposide for 1 h in 5% CO2 with humidified air, followed by three washes with TCM199, then cultured for 48 h in IVM medium with and without 10 µg/ml aphidicolin (Sigma, A0781). Fourth, etoposide followed by 2,3-dideoxycytidine (ddC; a nucleoside analogue) treatment. For this, freshly retrieved GV stage oocytes were cultured in IVM medium supplemented with 1 µg/ml etoposide for 1 h in 5% CO2 with humidified air, followed by three washes with TCM199, then cultured for 48 h in IVM medium with and without 10 μM ddc (SRL, 47248).

Washing experiment

Freshly retrieved GV stage oocytes were cultured in IVM medium supplemented with 1 µg/ml etoposide for 1 h in 5% CO2 with humidified air, followed by three washes with TCM199, then cultured for 24 h in IVM medium with and without 50 mM HU and then washed for 4 times with TCM199, followed by transfer to IVM medium and cultured up to 48 h.

Immunocytochemistry

Oocytes were fixed with 4% PFA, and processed for surface spreading of oocyte chromosomes as described previously (Qiao et al., 2018). Immunofluorescence staining was performed using the following primary antibodies against the following proteins with incubation overnight at room temperature: γH2AX (Millipore 05-636-I, 1:500), Pol δ (ImmunoTag, ITT1371, 1:100), Pol ε (ImmunoTag, ITT5065, 1:100), RFC1(ImmunoTag, ITT5314, 1:100), Pol beta (Elabscience, E-AB-31214, 1:100), PCNA (Cloud-Clone Crop PAA591Mi01, 1:100), RPA (Abcam, ab76420, 1:250), RAD51(Thermo Fisher Scientific, MA5-14419, 1:250). Slides were subsequently incubated with the following goat secondary antibodies for 1 h at 37°C: anti-rabbit-IgG conjugated to AF 488 (Thermo Fisher Scientific, 1:2000 dilution), anti-rabbit conjugated to AF 555 (Thermo Fisher Scientific, 1:2000), anti-mouse conjugated to AF 555 (Thermo Fisher Scientific, 1:1000), anti-mouse conjugated to AF 488 (Thermo Fisher Scientific, 1:1000). Coverslips were mounted with ProLong Dimond antifade reagent (Thermo Fisher Scientific).

Microinjection

Freshly retrieved GV stage oocytes were cultured in IVM medium followed by microinjections of PCNA, Pol δ, ε, Mad2, and negative control targeted siRNAs (Table S1) using a micromanipulator with thermal plate (Narishige, Japan) at 37°C. Then the microinjected oocytes were supplemented with 1 µg/ml etoposide for 1 h in 5% CO2 with humidified air, followed by three washes with TCM199, then cultured for 48 h in IVM medium.

Imaging and quantification

Immunofluorescence-stained oocytes and surface spreads were imaged using Zeiss Axio scope VII microscope with 10×, 40× Plan Apochromat 0.45 NA, or 63/100×Plan Apochromat 1.4 NA objectives and EXFO X-Cite metal halide light source. Images were captured with a Hamamatsu ORCA-ER CCD camera and processed using Zen software. All comparisons were made between datasets obtained from animals that were matched by age. Two observers performed all quantitative analyses; the second observer was blind to which group was being analyzed.

Statistical analysis

Data is presented as means±s.e.m. or s.d. and represents at least three independent experiments using 30 to 50 oocytes. Statistical tests, P and n values are described in the figure legends.

We thank the NIAB core microscope facility.

Author contributions

Conceptualization: A.K.S., S.L.K., R.B., A.M., B.K., H.B.D.P.R.; Methodology: A.K.S., S.L.K., R.B., A.M., B.K., H.B.D.P.R.; Validation: A.K.S., S.L.K., R.B., A.M., B.K., H.B.D.P.R.; Formal analysis: A.K.S., S.L.K., R.B., A.M., B.K., H.B.D.P.R.; Investigation: A.K.S., S.L.K., R.B., A.M., B.K., H.B.D.P.R.; Resources: H.B.D.P.R.; Data curation: H.B.D.P.R.; Writing - original draft: A.K.S., H.B.D.P.R.; Writing - review & editing: A.K.S., H.B.D.P.R.; Visualization: H.B.D.P.R.; Supervision: H.B.D.P.R.; Project administration: H.B.D.P.R.; Funding acquisition: H.B.D.P.R.

Funding

S.L.K. was supported by Department of Biotechnology, Ministry of Science and Technology, India (DBT) JRF. R.B. was supported by UGC JRF. A.M. was supported by CSIR JRF. This work was supported by a DBT grant (BT/PR31689/AAQ/1/747/2019) and Department of Science and Technology, Ministry of Science and Technology, India (DST) grants to H.B.D.P.R. (CRG/2018/002821), and DBT Ramalinga swami fellowship (BT/RLF/ Re-entry/21/2016) awarded to H.B.D.P.R..

The peer review history is available online at https://journals.biologists.com/jcs/article-lookup/doi/10.1242/jcs.257774.

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Competing interests

The authors declare no competing or financial interests.

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