Roscovitine and Trichostatin A promote DNA damage repair during porcine oocyte maturation

Bingyue Zhang A, Huiran NiuA, Qingqing Cai A, Mengqin Liao A, Keren Chen A, Yaosheng Chen A and Peiqing Cong A,B
AState Key Laboratory of Biocontrol, School of Life Sciences, Sun Yat-sen University, Guangzhou,
Guangdong, 510006, PR China.
BCorresponding author. Email: [email protected]

Abstract. Faithful repair of DNA double-strand breaks in mammalian oocytes is essential for meiotic maturation and embryonic development. In the present study we investigated the roles of Roscovitine and Trichostatin A (TSA) in DNA damage recovery during in vitro maturation of porcine oocytes. Etoposide was used to trigger DNA damage in oocytes. When these DNA-damaged oocytes were treated with 2 mM Roscovitine, 50 nM TSA or both for 22 h, first polar body extrusion and blastocyst formation in all treated groups were significantly improved compared with the etoposide-only group. The most significant improvement was observed when Roscovitine was present. Further immunofluorescent analysis of gH2A.X, an indicator of DNA damage, indicated that DNA damage was significantly decreased in all treated groups. This observation was further supported by analysing the relative mRNA abundance of DNA repair-related genes, including meiotic recombination 11 homolog A (MRE11A), breast cancer type 1 susceptibility protein (BRCA1), Recombinant DNA Repair Protein 51 (RAD51), DNA-dependent protein kinase catalytic subunit (PRKDC) and X-ray cross complementing gene 4 (XRCC4). Compared with the etoposide-only group, the experimental group with combined treatment of Roscovitine and TSA showed a significant decrease of all genes at germinal vesicle and MII stages. The Roscovitine-only treatment group revealed a similar tendency. Together, these results suggest that Roscovitine and TSA treatments could increase the capacity of oocytes to recover from DNA damage by enlisting DNA repair processes.

Additional keywords: CDK2, development, DSBs, HDAC.


Correct completion of oocyte meiosis ensures genomic integrity in mammals, which is essential for fertility and normal devel- opment. Meiosis initiates in fetal ovaries and is arrested at the dictyate stage of prophase I (defined as germinal vesicle stage, GV) before birth. This arrest persists until puberty, when mei- osis resumes regularly in a small subset of the oocyte population (Handel and Schimenti 2010). The protracted arrest of meiosis at prophase I makes oocytes highly sensitive to the accumulation of endogenous and environmentally induced damage, a com- mon example of which is DNA double-strand breaks (DSBs). Both endogenous metabolites or processes and exogenous fac- tors (e.g. UV, ionising radiation, physical hazards and chemical drugs) can induce DSBs (Ma et al. 2013; Halldorsson et al. 2016). There is evidence that female aging increases errors during chromosome segregation and promotes accumulation of DSBs in primordial follicles (Vijayalakshmi and Sharon 2008; Shiny et al. 2013). DNA damage in these cells can result in genetic abnormalities, which further lead to infertility, embry- onic death or developmental abnormalities (Marangos and Carroll 2012).

To protect oocyte development against threats posed by DSBs, cells have evolved mechanisms to detect DNA lesions, signal their presence and promote their repair (Jackson and Bartek 2009). In mammalian cells, there are two important pathways for repairing DSBs: the non-homologous end joining (NHEJ) pathway and the homologous recombination (HR) pathway. Another known pathway contributing to DSB repair, the single-strand annealing (SSA) pathway, is generally consid- ered as a sub-pathway of HR (Peter 2000). NHEJ is the major pathway of DSB repair in mammalian cells. Its core component is the Ku70–Ku80 DNA end-binding heterodimer, which binds the broken DNA end and activates the DNA-dependent protein kinase catalytic subunit (DNA-PKcs). DNA-dependent protein kinase (DNA-PK) is a serine–threonine protein kinase. It can induce conformational changes of the damage-recognition complex and interact with the ataxia-telangiectasia mutated (ATM) and ATM and Rad3-related protein (ATR). DNA-PKcs, ATM and ATR are the most important members of the phos- phoinositide-3-kinase-related protein kinase (PIKK) family in the response to DSBs (Huen and Chen 2008). Together, they phosphorylate p53, single-strand binding protein replication

Reproduction, Fertility and Development B. Zhang et al. protein A (RPA), Ku and X-ray cross complementing gene 4 (XRCC4). Subsequently, phosphorylated XRCC4 interacts with DNA ligase IV to form the DNA ligase IV (LIG4)–XRCC4 complex, which further interacts with XRCC4-like factor (XLF) to perform DNA ligation (Wang and Lees-Miller 2013; Stadler and Richly 2017).

The NHEJ pathway can directly ligate the break-ends of DNA without the need for a homologous template. When a sister template is available, however, the HR pathway becomes predominant. In this process, Recombinant DNA Repair Protein 51 (RAD51), a DNA strand exchange protein, is required to bind to the single-strand DNA (ssDNA; Ceccaldi et al. 2016). The loading of the RAD51 recombinase onto the ssDNA requires the breast cancer type 1 susceptibility protein (BRCA1), which also functions in the 5′ to 3′ resection of DSBs by colocalising with the resection complex MRE11 (Meiotic Recombination 11)– RAD50(Recombinant DNA Repair Protein 50)–NBS1 (Nijmegen Breakage Syndrome 1) (MRN) after DNA damage. The MRN complex is the main regulator of ATM activation. An important role for the ATM–ATR–DNA-PK kinase is to induce phosphorylation on serine 139 of the histone H2A variant, H2A. X (Jackson and Bartek 2009). Therefore, phosphorylation of H2A.X (gH2A.X) is frequently used as an indicator of DSB occurrence (Prakash et al. 2015; Stadler and Richly 2017).

The occurrence of DSB can trigger DNA repair processes. In most cancer studies, 2-(1-ethyl-2-hydroxyethylamino)- 6-benzylamino-9-isopropylpurine (Roscovitine) has been reported to hinder DNA repair, either by inhibiting the NHEJ pathway or by inducing DNA damage signalling genes P53– RAD51–BRCA1 and promoting apoptosis (Deans et al. 2006; Federico et al. 2010; Wang and Kim 2016). Nonetheless, the recovery from DNA damage in some cancer cells after Roscovitine treatment has also been reported (Crescenzi et al. 2005). As a highly efficient and selective inhibitor of cyclin- dependent kinases (CDKs), Roscovitine mainly targets CDK2, but can also affect CDKs4/6 (Meijer et al. 1997). Since CDKs play an essential role in the intracellular control of the cell division cycle (CDC), Roscovitine provides a useful tool for cell cycle studies, especially those related to cancer treatments.

In cancer cells, histone acetylation, which is regulated by histone acetyl transferases (HATs) and histone deacetylases (HDACs), may also play a role in DNA damage and repair. HDAC inhibitors have been reported to abolish drug-resistance, to sensitise cells to chemotherapy, to decrease NHEJ activity and to affect the expression of BRCA1, KU70, KU80, DNA- PKcs and the activity of KU70 (Bohrer et al. 2014; Jin et al. 2014; Robert et al. 2016; Roos and Krumm 2016). Inappropriate levels of histone acetylation also result in compromised oocyte meiosis and maturation. There is evidence that HDAC inhibition can improve embryonic development, reduce DSBs and increase the cleavage and blastocyst rates in early porcine embryos (Bohrer et al. 2014; Wang et al. 2015a). As an inhibitor of histone deacetylases, Trichostatin A (TSA) has been reported to facilitate epigenetic reprogramming during the development of cloned animals (Ji et al. 2013). The beneficial effect of TSA on oocyte in vitro maturation is, however, mainly ascribed to the prolonged maturation process and the delayed aging of oocytes (Jesˇeta et al. 2008; Jin et al. 2014). Since few studies have examined the roles of Roscovitine and TSA in DNA damage response in porcine oocytes, this study aimed to see if these drugs could help porcine oocytes to recover from DNA double- strand breaks during in vitro maturation.

Materials and methods

Porcine oocyte collection and in vitro maturation (IVM)

Porcine ovaries were collected from a local abattoir and trans- ported to the laboratory in 0.9% (w/v) saline with antibiotics at 30–358C within 3 h. Antral follicles (2 to 6 mm in diameter) were aspirated with a 16-gauge needle connected to a 10-mL syringe. Follicular fluid-containing cumulus–oocyte complexes (COCs) were resuspended with Hepes-buffered Tyrode’s lactate medium containing 0.1% (w/v) polyvinyl alcohol (TL–PVA) and observed under a stereomicroscope. Oocytes with evenly granulated cytoplasm and at least three uniform layers of cumulus cells were selected, washed three times with tissue culture medium 199 (TCM199; Sigma) plus 5% (v/v) fetal bovine serum (FBS) and cultured at 38.58C in an atmosphere of 5% CO2 in humidified air in 12-well plates (Thermo) containing 1 mL maturation medium. The maturation medium was a TCM199-based medium plus 0.57 mM L-cysteine, 10% (v/v) porcine follicular fluid (PFF), 0.5 mg mL—1 follicle-stimulating hormone (FSH), 0.5 mg mL—1 luteinising hormone (LH) and 10 ng mL—1 epidermal growth factor (EGF). After 22 h of maturation, COCs were transferred to the maturation medium without L-cysteine and hormones for another 22 h (Cong et al. 2008; Kong et al. 2014).

Drug treatments

Etoposide, Roscovitine and Trichostatin A (all from Selleck) were dissolved in dimethyl sulphoxide (DMSO). Roscovitine (2 mM) and–or TSA (50 nM) was added into the 0–22 h matu- ration medium. Etoposide was used for 5 h to trigger DNA damage in premature porcine oocytes immediately after IVM started, which means for the first 5 h oocytes were cultured in 0–22 h maturation medium without Etoposide (control) or with Etoposide (Etp) or with both Etoposide and Roscovitine/TSA (drug treatment groups). Then oocytes were moved into the exact same culture media without Etoposide and cultured for another 17 h before being moved to 22–44 h maturation medium.

Preparation of denuded oocytes

COCs at desired stages of maturation were collected and cumulus cells were removed by vigorous pipetting in the manipulation medium with 0.1% hyaluronidase. Those denuded oocytes were collected for subsequent experiments (Cong et al. 2008).

Observation of polar body extrusion

After maturation for 44 h, denuded oocytes were transferred to the manipulation medium supplemented with 5% (w/v) sucrose and incubated for 30 min before observation of polar body extrusion. A glass needle was used to manipulate oocytes for an all-round observation. For calculation of percentages of polar body extrusion, each treatment was repeated 4–6 times with each replicate containing ,50 oocytes.

Parthenogenetic activation and embryo culture

Oocytes with the first polar body were activated with a single DC pulse of 120 kV cm—1 for 30 ms using an Electro Cell Manipulator ECM-2001 (BTX) in the fusion solution. After washing twice in the embryo culture medium PZM-5, the activated oocytes were cultured in 4-well plates for 7–9 days to observe cleavage and blastocyst formation. For calculation of cleavage rates and blastocyst rates, each treatment was repeated 4–6 times with each replicate containing ,50 oocytes.

Immunofluorescence analysis

Oocytes were fixed with 4% paraformaldehyde in phosphate- buffered saline (PBS) for 45 min at room temperature, followed by permeabilising in 0.5% Triton X-100 in PBS for 30 min. After three washes in PBS containing 0.1% Tween 20 and 0.01% Triton X-100, oocytes were blocked in 1% bovine serum albu- min (BSA) in PBS for 1 h at room temperature and then incu- bated overnight at 48C with phospho-histoneH2A.X (Ser139) rabbit mAb (Cell Signalling Technology; diluted 1 : 400 in 5% BSA in PBS). After three more washes, oocytes were labelled with Chromeo 488 Alkyne (Active Motif; diluted 1 : 1 000 in 5% BSA in PBS) for 1–2 h at room temperature, immediately followed by another three washes. Oocyte DNA was then stained by 4′,6-diamidino-2-phenylindole (DAPI; Sigma; diluted in PBS) for 20–30 min at room temperature. After four more washes, oocytes were laid on glass slides with Vectashield (Vector). Samples were observed and photographed under a fluorescent microscope (AxioImgaerZ1; Carl Zeiss). Camera settings (including exposure time) were the same in all obser- vations. Each experiment group consisted of at least 20 oocytes and each treatment was repeated 3–4 times.

Real-time quantitative polymerase chain reaction (PCR)

Total RNA was extracted from ,100 cumulus-free oocytes from each group (including control, Etoposide, Roscovitine and–or TSA treatment groups) using a commercial RNA isolation kit (RNAprep Pure Micro Kit; Tiangen), including a DNase I solu- tion treatment for 15 min for each sample. This was followed by reverse transcription using HisScript II Q RT SuperMix for Qpcr ( gDNA wiper; Vazyme) according to the manufacturer’s instructions: 14 mL of each RNA sample was mixed with 8 mL 4 gDNA wiper Mix and 10 mL RNase-free double distilled water (ddH2O) in a 1.5-mL reaction tube and the mixture was incubated in a 428C water bath for 2 min, then 8 mL 5 HiScript II qRT SuperMix II was added into the mixture and a total volume of 40 mL mixture was incubated in a 508C water bath for 15 min, then 858C for 5 s and then cooled on ice. Samples were frozen at 808C for storage and thawed for experiment within a week.

A 10 mL reaction volume included 0.5 mL cDNA sample, 5 mL Hieff qPCR 2 SYBR Green Master Mix (Yeasen), 3.5 mL RNase-free water and 0.5 mL each of forward and reverse gene- specific primers. Thereafter, quantitative real-time PCR was performed using the LightCycler 480 (Roche). Cycling condi- tions were composed of denaturation at 958C for 5 min, followed by 40 three-step cycles of 958C for 10 s, 588C for 20 s and 728C for 20 s, then a melting cycle of 958C for 15 s, starting at 608C with a 0.28C s—1 transition rate to 958C. Each sample was run in quadruplicate. Cycle quantification (Cq) of the no template control (NTC) is 35. Primer pairs were designed by Primer Premier 6.0 (Premier Biosoft) and checked using Primer-Blast software (National Center for Biotechnology Information). Primer sequences and the approximate sizes of the amplified fragments for all transcripts are listed in Table 1. To determine specificity of the reaction, PCR products were analysed by gel electrophoresis and dissociation curve analysis. Glyceralde- hyde-3-phosphate dehydrogenase (GAPDH) was used as an internal standard. All experiments contained three biological replicates. Results of real-time PCR were calculated by the DDCT method to compare relative abundances of target genes.

Statistical analysis

Comparisons between multiple groups within a single time point were performed by one-way analysis of variance (ANOVA) followed by Dunnett’s test. Normality and homogeneity of variances of data were confirmed before running ANOVA. As for comparing the overall results of gH2A.X immunofluores- cence and gene expression levels, a mixed model was per- formed. Two-tailed P , 0.05 was considered to be significant, whereas 0.05 # P , 0.1 was considered to indicate tendencies. Relative fluorescence intensity of gH2A.X was calculated by Image Pro Plus 6.0 (Media Cybernetics Inc.). Graphs were created using Prism 5 (GraphPad Software Inc.) and all quan- titative data are presented as mean s.e.m. SPSS 23.0 (IBM) was used for all data analysis.

Fig. 1. Foundation of the DNA-damage model in porcine oocytes. (a–d”’) Immunofluorescence of porcine oocytes incubated with (a–a”’) 0, (b–b”’) 100, (c–c”’) 300 or (d–d”’) 500 mg mL—1 etoposide for 5 h. The degree of DNA damage was assessed by staining with the gH2A.X antibody (green). DNA was stained with DAPI (blue). Scale bar = 50 mm. (e) gH2A.X intensity was quantified for each group. Data are presented as mean s.e.m; n , 90 each group. ***P , 0.001.


Building of a DNA-damage model in porcine oocytes by etoposide treatment

In order to build a DNA-damage model in oocytes and deter- mine the optimal concentration of etoposide, oocytes collected from ovaries were exposed to 100, 300 or 500 mg mL—1 etopo- side for 5 h from the beginning of IVM. They were then subjected to the assessment of DSBs by examining the relative labelling intensity of gH2A.X. As shown in Fig. 1, etoposide treatment led to a significant, dose-dependent increase in gH2A. X labelling (Fig. 1b’–e), while it remained at low basal levels in populations of untreated control oocytes (Fig. 1a’). Since the
degree of DNA damage (as indicated by gH2A.X labelling) reached 2-fold that of the control group at 300 mg mL—1 whilst maintaining a reasonable embryo survival rate (Fig. 1e), 300 mg mL—1 etoposide was chosen as the optimal concentration for establishing the DSB model in oocytes.

Effects of treatment with Roscovitine and–or TSA on meiotic and embryonic development of DNA-damaged oocytes

To test the effects of Roscovitine and–or TSA, the etoposide- only group was used as a control. Etoposide-treated oocytes were then incubated with 2 mM Roscovitine and–or 50 nM TSA for 22 h. The rates of the first polar body extrusion (PBE) were examined after maturation. The results are shown in Fig. 2a. When treated with etoposide only, the PBE rate of the oocytes was significantly lower than that in the non-treatment group (42.82% vs 60.37%), suggesting that DSB induced by etoposide prevented meiotic progress of the oocytes. Roscovitine treat- ment significantly increased the PBE rate of etoposide-treated oocytes up to 58.58% (P 0.001), which was similar to the combined effect of Roscovitine and TSA (58.51%; P 0.006). However, although TSA treatment also increased the PBE rate (52.84%) in etoposide-treated oocytes, its effect was not sig- nificant (P 0.069; Fig. 2a). The concentrations chosen for Roscovitine and TSA were selected via concentration effectivity experiments, the results of which are shown in Figs S1–S3, available as Supplementary Material to this paper.

Fig. 2. Effects of TSA and–or Roscovitine treatment during IVM on nuclear maturation and embryonic development after parthenogenesis. (a) The rates of the first polar body extrusion. (b) The cleavage rates. (c) The blastocyst rates. Data are presented as mean s.e.m. Columns with different letters differ significantly; n , 240 each group. P , 0.05.

To further assess the developmental capacity of oocytes, parthenogenetic activation was carried out after IVM and the cleavage rate and blastocyst formation were evaluated (Fig. 2b, c). There were no significant differences in cleavage rates between groups regardless of treatment (Fig. 2b). How- ever, etoposide treatment significantly reduced blastocyst for- mation from 20.48% to 5.00% (P , 0.001). Roscovitine treatment and the combined treatment of Roscovitine and TSA recovered the rates to 12.67% (P , 0.05) and 17.30% (P , 0.05) respectively, which were significantly higher than for the etoposide-only group. No significant difference in blastocyst formation was observed between the TSA and the etoposide-only group (Fig. 2c).

Effects of Roscovitine and–or TSA treatment on DNA damage during IVM

To understand the above results, we then examined DNA damage in all groups by immunofluorescent labelling of the gH2A.X, the presence of which is induced by DNA damage response kinases ATM, ATR or DNA-PK in the vicinity of damage sites (Yuan et al. 2010). The examination was per- formed at four different time points (5, 24, 36, 44 h), repre- senting the GV, germinal vesicle breakdown (GVBD), MI and MII stages of meiosis respectively. The fluorescence intensities of gH2A.X of all groups after 24 h of maturation are shown in Fig. 3. The labelling of gH2A.X was almost invisible in the non-treatment group (Fig. 3a’), but was clearly observed in all etoposide-treated groups (Fig. 3b’–e’). However, when treated with Roscovitine and–or TSA, the intensity of gH2A.X was significantly reduced at all time points (Fig. 3c’, d’, e’, f and Figs S4–S6), with the Roscovitine group showing the most signifi- cant drop. Notably, the gH2A.X intensity continued to decrease
over time in these groups, which may reflect the fact that the self-repair mechanism in the oocyte operates during in vitro maturation. Together, these observations suggested that TSA and Roscovitine, especially the latter, could promote DNA damage repair in DNA-damaged oocytes, leading to improved PBE and blastocyst formation.

Expression of DNA repair-related genes at different stages of oocyte meiotic maturation

To further confirm the influence of Roscovitine and TSA on DNA damage recovery, the expression of DNA repair-related genes (MRE11A, BRCA1, RAD51, DNA-dependent protein kinase catalytic subunit (PRKDC) and XRCC4) was then examined in GV, GVBD, MI and MII oocytes (Fig. 4). The expression of all genes followed a similar overall trend; the gene transcripts accumulated through GV stage, reached a relatively high level at GVBD, decreased through MI stage and accrued again until MII. Treatment with etoposide, Roscovitine and TSA did not affect this gene expression trend throughout meiosis. However, compared with the control, etoposide significantly downregulated all the genes at GV but upregulated them at GVBD and MI and induced a dramatic rise at MII. When DNA- damaged oocytes were treated with TSA, the expression levels of MRE11A, BRCA1and PRKDC increased until 36 h (Fig. 4a, b, d), whereas RAD51 and XRCC4 were upregulated only at 24 h (Fig. 4c, e). The levels of MRE11A were significantly reduced by Roscovitine treatment at the initial and final time points. Other genes in this group were affected similarly but to a lesser extent. When co-treated with both Roscovitine and TSA, all the genes except XRCC4 were increased at 24 h (the same as the TSA treatment group), but only MRE11A was upregulated at 36 h. However, compared with the etoposide-only group, all the genes displayed a significant decrease at 5 h and 44 h after combined treatment with Roscovitine and TSA (Fig. 4a–e).


The present study aimed to unravel the potentially beneficial effects of Roscovitine and TSA on DNA damage recovery during in vitro maturation of porcine oocytes. To this end, we first built a DNA-damage model in porcine oocytes by using etoposide (Fig. 1). Although cleavage rates didn’t show large differences, the PBE rates and blastocyst rates were significantly increased, suggesting that the developmental competence of these DNA-damaged oocytes was significantly improved when treated with 2 mM Roscovitine and–or 50 nM TSA (Fig. 2). We suspect that those oocytes that successfully managed to transi- tion through meiosis with the different treatments had similar chances to start a division but the correct proportions were different. This is likely to be due to the reduction of DNA damage (as demonstrated by gH2A.X labelling) throughout the process of meiosis after these treatments (Fig. 3 and Figs S1–S3).

Fig. 3. Effects of TSA and–or Roscovitine treatment on DNA damage in porcine oocytes. Oocytes with or without drug treatment as described in ‘Materials and methods’ were matured in vitro for 5, 24, 36 or 44 h. (a–e”) Immunofluorescence of the samples matured for 24 h. The degree of DNA damage was assessed by staining with the gH2A.X antibody (green) and DNA was stained with DAPI (blue). (a–a”) Con, control oocytes, (b–b”) Etp, etoposide-only group, (c–c”) E+TSA, etoposide and 50 nM TSA co-treated group, (d–d”) E+Ros, etoposide and 2 mM Roscovitine co-treated group, (e–e”) E+TSA+Ros, etoposide, 50 nM TSA and 2 mM Roscovitine co-treated group. Scale bar = 50 mm; n , 90 each group (f) gH2A.X intensity analysis in each group. Data are presented as mean s.e.m. Comparisons between control and etoposide treatment are indicated by ###P , 0.001. Comparison of etoposide treatment group and the other groups are indicated by ***P , 0.001.

In addition, despite similar expression trends throughout the process of meiosis, DNA repair-related genes showed signifi- cantly different levels of expression between treatment groups.

Fig. 4. Relative mRNA expression of (a) MRE11A, (b) BRCA1, (c) RAD51, (d) PRKDC and (e) XRCC4 at four meiotic stages during porcine oocyte maturation. The mRNA abundance of all the genes was normalised to that of GAPDH and the graph shows relative gene expression in each group. Con, control oocytes; Etp, etoposide-only group; E+TSA, etoposide and 50 nM TSA co-treated group; E+Ros, etoposide and 2 mM Roscovitine co-treated group; E TSA Ros, etoposide, 50 nM TSA and 2 mM Roscovitine co-treated group; n 270 each group. Comparisons between control and etoposide treatment are indicated by ###P , 0.001; ##P , 0.01; #P , 0.05. Comparison of etoposide treatment group and the other groups are indicated by ***P , 0.001; **P , 0.01; *P , 0.05.

The genes that we chose are either implicated in the non- homologous end joining (NHEJ) pathway or involved in the homologous recombination (HR) pathway. In the HR pathway, DNA damage is sensed by the MRN complex (consisting of MRE11, RAD50 and NBS1), which co-localises with BRCA1. BRCA1 helps to load RAD51 to the ssDNA which is needed as a template (Oktay et al. 2015). In the NHEJ pathway, PRKDC is the gene encoding DNA-PKcs, which induces conformational changes of the damage recognition complex and phosphorylates XRCC4, an essential participant in the religation of the DNA ends. These genes were expressed at low levels at 5 h (Fig. 4), when oocytes were still exposed to etoposide-treatment. The etoposide in the maturation medium consecutively causes DSBs and obstructs DNA repair. After 5 h, etoposide was removed from the medium. The transcript abundance of these genes in DNA-damaged oocytes then increased and peaked at 44 h (Fig. 4), suggesting that MII is an important stage for DNA repair. Compared with the etoposide-only group, MRE11A showed a significant decrease at 5 h and 44 h in the Roscovitine treatment group; however, in the TSA treatment group, it was significantly upregulated at 5 h, 24 h and 36 h. The group treated with both Roscovitine and TSA seemed to experience a com- bined effect, with Roscovitine predominating at 5 h and 44 h. Despite the difference in expression levels, all the other genes showed similar trends of expression after experimental treat- ments.

Notably, all genes, with no exception, were significantly decreased at 5 h and 44 h in the Roscovitine–TSA combined treatment group. Taking into account the results of oocyte development and gH2A.X labelling, it is reasonable to suggest that these drugs have their maximum influence at the GV and MII stages, while the treatment is effective throughout the process of meiosis. These two stages may represent two key time points for DNA damage repair during meiosis of the oocyte.

High doses of Roscovitine were previously reported to induce apoptosis in tumour cells (Mgbonyebi et al. 1999; Hahntow et al. 2004). Although Roscovitine has been widely investigated in studies of combination chemotherapy for cancer (Abaza et al. 2008; Appleyard et al. 2009), its effect on DSBs is still not well understood. The effects of Roscovitine may vary in different cell types. For example, in HCT116 and H1299 cell lines, combined treatments with doxorubicin and Roscovitine increased the frequency of double-strand breaks and sensitised cells to doxorubicin (Crescenzi et al. 2005). In other cell lines, such as A549 and HEC1B, however, it can decrease double- strand breaks and enhance clonogenic survival (Crescenzi et al. 2005). There are studies reporting that Roscovitine could modulate proteins involved in the process of DNA damage response by CDK inhibition (Maude and Enders 2005; Tian et al. 2009). Another study found that Roscovitine reduces the extent of DNA damage mediated by Camptothecin and suggests a novel mechanism of apoptosis prevention (Pizarro et al. 2011). The main target of Roscovitine, CDK2, has been proven to interact with MRE11 and participate in double-strand break repair (Buis et al. 2012).

Previous research on the influence of Roscovitine treatment on porcine oocytes focussed on meiotic arrest maintenance and indicated that a high dose of Roscovitine could help to improve oocyte developmental competence by effectively blocking meiosis in the absence of LH and by promoting oocyte cytoplasmic maturation (Zhang et al. 2017). In our research, we found that Roscovitine could reduce DSBs in DNA-damaged porcine oocytes and modulate DNA repair-related genes. Therefore, our study argues for a role of Roscovitine in the DNA damage response in germ cells.

Histone acetylation, which can be regulated by HDACs and is involved in regulating chromatin dynamics, may also influence the DNA damage response (Santos-Rosa et al. 2002; Choudhary et al. 2009). Therefore, nonselective HDAC inhibi- tors are widely tested for use in anticancer combination thera- pies (Nolan et al. 2008; Poljakova et al. 2011; Groselj et al. 2013). HDAC inhibition has been found to exacerbate cytotox- icity of DSB-inducing anticancer drugs, suppress the DNA damage response and result in more DSBs in cortical neurons, yeast strains and human lung cancer cells (Wang et al. 2012, 2015b; Vashishta and Hetman 2014). However, it is not known if HDAC inhibition is correlated with DNA damage responses in porcine oocytes. There is evidence that HDAC inhibition improves the efficiency of DSB repair in porcine preimplanta- tion embryos (Wang et al. 2015a). We postulated that as a nonselective HDAC inhibitor, TSA might also affect DSB recovery in porcine oocytes and the combined treatment with both Roscovitine and TSA might have an even more dramatic effect. This hypothesis was supported by our observation that Roscovitine and–or TSA treatments significantly reduced the levels of DNA damage in oocytes (Fig. 2), with the most significant reduction being observed in the Roscovitine treat- ment groups (Roscovitine alone group and Roscovitine–TSA group). However, as long as Roscovitine was present, no significant difference was observed between these groups, suggesting that Roscovitine contributed more to DNA damage recovery than TSA.

In conclusion, our findings provide two major insights into the effects of Roscovitine and TSA on DNA damage repair during in vitro maturation of porcine oocytes. Firstly, treatment with TSA or Roscovitine or both could reduce the degree of DNA damage, but only when the reduction is sufficient could a beneficial effect on the developmental potential of the oocytes be obtained. In our study, the combined treatment with both drugs rescued PBE and blastocyst formation almost back to normal levels. Secondly, the mRNA abundance of DNA repair- related genes demonstrated a significant reduction at GV and MII, indicating that these are the most important stages of DNA damage recovery during meiosis.

Conflicts of interest

The authors declare no conflicts of interest to this work.


This work was supported by NSFC-Guangdong Joint Fund (U1201213), National Transgenic Breeding Program (2016ZX08006003–006) and Science And Technology Project of Guangdong Province (2014B020202001).


Abaza, M. S. I., Bahman, A.-M. A., and Al-Attiyah, R. J. (2008). Roscovitine synergizes with conventional chemo-therapeutic drugs to induce effi- cient apoptosis of human colorectal cancer cells. World J. Gastroenterol. 14, 5162–5175. doi:10.3748/WJG.14.5162
Appleyard, M. V., O’Neill, M. A., Murray, K. E., Paulin, F. E., Bray, S. E., Kernohan, N. M., Levison, D. A., Lane, D. P., and Thompson, A. M. (2009). Seliciclib (CYC202, R-roscovitine) enhances the antitumor effect of doxorubicin in vivo in a breast cancer xenograft model. Int. J. Cancer 124, 465–472. doi:10.1002/IJC.23938
Bohrer, R. C., Duggavathi, R., and Bordignon, V. (2014). Inhibition of histone deacetylases enhances DNA damage repair in SCNT embryos. Cell Cycle 13, 2138–2148. doi:10.4161/CC.29215
Buis, J., Stoneham, T., Spehalski, E., and Ferguson, D. O. (2012). Mre11 regulates CtIP-dependent double-strand break repair by interac- tion with CDK2. Nat. Struct. Mol. Biol. 19, 246–252. doi:10.1038/ NSMB.2212
Ceccaldi, R., Rondinelli, B., and D’Andrea, A. D. (2016). Repair pathway choices and consequences at the double-strand break. Trends Cell Biol. 26, 52–64. doi:10.1016/J.TCB.2015.07.009
Choudhary, C., Kumar, C., Gnad, F., Nielsen, M. L., Rehman, M., Walther,
T. C., Olsen, J. V., and Mann, M. (2009). Lysine acetylation targets protein complexes and co-regulates major cellular functions. Science 325, 834. doi:10.1126/SCIENCE.1175371
Cong, P. Q., Kim, E. S., Song, E. S., Yi, Y. J., and Park, C. S. (2008). Effects of fusion/activation methods on development of embryos produced by nuclear transfer of porcine fetal fibroblast. Anim. Reprod. Sci. 103, 304–
311. doi:10.1016/J.ANIREPROSCI.2006.12.012
Crescenzi, E., Palumbo, G., and Brady, H. J. (2005). Roscovitine modulates DNA repair and senescence: implications for combination chemother- apy. Clin. Cancer Res. 11, 8158–8171. doi:10.1158/1078-0432.CCR- 05-1042
Deans, A. J., Khanna, K. K., McNees, C. J., Mercurio, C., Heierhorst, J., and McArthur, G. A. (2006). Cyclin-dependent kinase 2 functions in normal DNA repair and is a therapeutic target in BRCA1-deficient cancers. Cancer Res. 66, 8219–8226. doi:10.1158/0008-5472.CAN-05-3945
Federico, M., Symonds, C. E., Bagella, L., Rizzolio, F., Fanale, D., Russo, A., and Giordano, A. (2010). R-Roscovitine (Seliciclib) prevents DNA damage-induced cyclin A1 upregulation and hinders non-homologous end-joining (NHEJ) DNA repair. Mol. Cancer 9, 208. doi:10.1186/1476- 4598-9-208
Groselj, B., Sharma, N. L., Hamdy, F. C., Kerr, M., and Kiltie, A. E. (2013). Histone deacetylase inhibitors as radiosensitisers: effects on DNA damage signalling and repair. Br. J. Cancer 108, 748–754. doi:10. 1038/BJC.2013.21
Hahntow, I. N., Schneller, F., Oelsner, M., Weick, K., Ringshausen, I., Fend, F., Peschel, C., and Decker, T. (2004). Cyclin-dependent kinase inhibitor Roscovitine induces apoptosis in chronic lymphocytic leukemia cells. Leukemia 18, 747–755. doi:10.1038/SJ.LEU.2403295
Halldorsson, B. V., Hardarson, M. T., Kehr, B., Styrkarsdottir, U., Gylfason, A., Thorleifsson, G., Zink, F., Jonasdottir, A., Jonasdottir, A., Sulem, P., Masson, G., Thorsteinsdottir, U., Helgason, A., Kong, A., Gudbjartsson,
D. F., and Stefansson, K. (2016). The rate of meiotic gene conversion varies by sex and age. Nat. Genet. 48, 1377–1384. doi:10.1038/NG.3669 Handel, M. A., and Schimenti, J. C. (2010). Genetics of mammalian meiosis: regulation, dynamics and impact on fertility. Nat. Rev. Genet. 11, 124–
136. doi:10.1038/NRG2723
Huen, M. S., and Chen, J. (2008). The DNA damage response pathways: at the crossroad of protein modifications. Cell Res. 18, 8–16. doi:10.1038/ CR.2007.109
Jackson, S. P., and Bartek, J. (2009). The DNA-damage response in human biology and disease. Nature 461, 1071–1078. doi:10.1038/ NATURE08467 Jesˇeta, M., Petr, J., Krejcˇova´, T., Chmel´ıkova´, E., and J´ılek, F. (2008). In vitro ageing of pig oocytes: effects of the histone deacetylase inhibitor trichostatin A. Zygote 16, 145–152. doi:10.1017/S0967199408004668
Ji, Q., Zhu, K., Liu, Z., Song, Z., Huang, Y., Zhao, H., Chen, Y., He, Z., Mo, D., and Cong, P. (2013). Improvement of porcine cloning efficiency by trichostain A through early-stage induction of embryo apoptosis. Therio- genology 79, 815–823. doi:10.1016/J.THERIOGENOLOGY.2012.12.010
Jin, Y. X., Zhao, M. H., Zheng, Z., Kwon, J. S., Lee, S. K., Cui, X. S., and Kim, N. H. (2014). Histone deacetylase inhibitor trichostatin A affects porcine oocyte maturation in vitro. Reprod. Fertil. Dev. 26, 806–816. doi:10.1071/RD13013
Kong, Q., Xie, B., Li, J., Huan, Y., Huang, T., Wei, R., Lv, J., Liu, S., and Liu, Z. (2014). Identification and characterization of an oocyte factor required for porcine nuclear reprogramming. J. Biol. Chem. 289, 6960– 6968. doi:10.1074/JBC.M113.543793
Ma, J.-Y., Ou-Yang, Y.-C., Wang, Z.-W., Wang, Z., Jiang, Z.-Z., Luo,
S.-M., Hou, Y., Liu, Z., Schatten, H., and Sun, Q.-Y. (2013). The effects of DNA double-strand breaks on mouse oocyte meiotic maturation. Cell Cycle 12, 1233–1241. doi:10.4161/CC.24311
Marangos, P., and Carroll, J. (2012). Oocytes progress beyond prophase in the presence of DNA damage. Curr. Biol. 22, 989–994. doi:10.1016/ J.CUB.2012.03.063
Maude, S. L., and Enders, G. H. (2005). CDK inhibition in human cells compromises Chk1 function and activates a DNA damage response. Cancer Res. 65, 780.
Meijer, L., Borgne, A., Mulner, O., Chong, J. P. J., Blow, J. J., Inagaki, N., Inagaki, M., Delcros, J.-G., and Moulinoux, J.-P. (1997). Biochemical and cellular effects of Roscovitine, a potent and selective inhibitor of the cyclin-dependent kinases cdk2, cdk2 and cdk5. Eur. J. Biochem. 243, 527–536. doi:10.1111/J.1432-1033.1997.T01-2-00527.X
Mgbonyebi, O. P., Russo, J., and Russo, I. H. (1999). Roscovitine induces cell death and morphological changes indicative of apoptosis in MDA- MB-231 breast cancer cells. Cancer Res. 59, 1903.
Nolan, L., Johnson, P. W., Ganesan, A., Packham, G., and Crabb, S. J. (2008). Will histone deacetylase inhibitors require combination with other agents to fulfil their therapeutic potential? Br. J. Cancer 99, 689– 694. doi:10.1038/SJ.BJC.6604557
Oktay, K., Turan, V., Titus, S., Stobezki, R., and Liu, L. (2015). BRCA mutations, DNA repair deficiency, and ovarian aging. Biol. Reprod. 93, 67. doi:10.1095/BIOLREPROD.115.132290
Parran, P. (2000). DNA double strand break repair in mammalian cells. Curr. Opin. Genet. Dev. 10, 144–150. doi:10.1016/S0959-437X(00)00069-1
Pizarro, J. G., Folch, J., Junyent, F., Verdaguer, E., Auladell, C., Beas- Zarate, C., Pallas, M., and Camins, A. (2011). Antiapoptotic effects of Roscovitine on camptothecin-induced DNA damage in neuroblastoma cells. Apoptosis 16, 536–550. doi:10.1007/S10495-011-0583-3
Poljakova, J., Hrebackova, J., Dvorakova, M., Moserova, M., Eckschlager, T., Hrabeta, J., Go¨ttlicherova, M., Kopejtkova, B., Frei, E., and Kizek, R. (2011). Anticancer agent ellipticine combined with histone deacetylase inhibitors, valproic acid and trichostatin A, is an effective DNA damage strategy in human neuroblastoma. Neuroendocrinol. Lett. 32, 101.
Prakash, R., Zhang, Y., Feng, W., and Jasin, M. (2015). Homologous recombination and human health: the roles of BRCA1, BRCA2, and associated proteins. Cold Spring Harb. Perspect. Biol. 7, a016600. doi:10.1101/CSHPERSPECT.A016600
Robert, C., Nagaria, P. K., Pawar, N., Adewuyi, A., Gojo, I., Meyers, D. J., Cole, P. A., and Rassool, F. V. (2016). Histone deacetylase inhibitors decrease NHEJ both by acetylation of repair factors and trapping of PARP1 at DNA double-strand breaks in chromatin. Leuk. Res. 45, 14–23. doi:10.1016/J.LEUKRES.2016.03.007
Roos, W. P., and Krumm, A. (2016). The multifaceted influence of histone deacetylases on DNA damage signalling and DNA repair. Nucleic Acids Res. 44, 10017–10030.
Santos-Rosa, H., Schneider, R., Bannister, A. J., Sherriff, J., Bernstein, B. E., Emre, N. C., Schreiber, S. L., Mellor, J., and Kouzarides, T. (2002). Active genes are tri-methylated at K4 of histone H3. Nature 419, 407–
411. doi:10.1038/NATURE01080
Shiny, T., Fang, L., Robert, S., Komala, A., Evrim, U., Kyungah, J., Maura, D., Mark, R., Fred, M., Sumanta, G., and Kutluk, O. (2013). Impairment of BRCA1-related DNA double strand break repair leads to ovarian aging in mice and humans. Sci Transl Med. 13, 172–221. doi:10.1126/ SCITRANSLMED.3004925
Stadler, J., and Richly, H. (2017). Regulation of DNA repair mechanisms: how the chromatin environment regulates the DNA damage response. Int. J. Mol. Sci. 18, E1715. doi:10.3390/IJMS18081715
Subramanian, V. V., and Bickel, S. E. (2008). Aging predisposes oocytes to meiotic nondisjunction when the cohesin subunit Smc1 is reduced. PLoS Genet. 14, e1000263. doi:10.1371/JOURNAL.PGEN.1000263
Tian, B., Yang, Q., and Mao, Z. (2009). Phosphorylation of ATM by Cdk5 mediates DNA damage signalling and regulates neuronal death. Nat. Cell Biol. 11, 211–218. doi:10.1038/NCB1829
Vashishta, A., and Hetman, M. (2014). Inhibitors of histone deacetylases enhance neurotoxicity of DNA damage. Neuromolecular Med. 16, 727– 741. doi:10.1007/S12017-014-8322-X
Wang, H., and Kim, N. H. (2016). CDK2 is required for the DNA damage response during porcine early embryonic development. Biol. Reprod. 95, 31. doi:10.1095/BIOLREPROD.116.140244
Wang, C., and Lees-Miller, S. P. (2013). Detection and repair of ionizing radiation-induced DNA double strand breaks: new developments in nonhomologous end joining. Int. J. Radiat. Oncol. Biol. Phys. 86, 440–449. doi:10.1016/J.IJROBP.2013.01.011
Wang, H., Zhou, W., Zheng, Z., Zhang, P., Tu, B., He, Q., and Zhu, W. G. (2012). The HDAC inhibitor depsipeptide transactivates the p53/p21 pathway by inducing DNA damage. DNA Repair (Amst.) 11, 146–156. doi:10.1016/J.DNAREP.2011.10.014
Wang, H., Luo, Y., Lin, Z., Lee, I. W., Kwon, J., Cui, X. S., and Kim, N. H. (2015a). Effect of ATM and HDAC inhibition on etoposide-induced DNA damage in porcine early preimplantation embryos. PLoS One 10, e0142561. doi:10.1371/JOURNAL.PONE.0142561
Wang, S. H., Lin, P. Y., Chiu, Y. C., Huang, J. S., Kuo, Y. T., Wu, J. C., and Chen, C. C. (2015b). Curcumin-mediated HDAC inhibition suppresses the DNA damage response and contributes to increased DNA damage sensitivity. PLoS One 10, e0134110. doi:10.1371/JOURNAL.PONE. 0134110
Yuan, J., Adamski, R., and Chen, J. (2010). Focus on histone variant H2AX: to be or not to be. FEBS Lett. 584, 3717–3724. doi:10.1016/J.FEBSLET. 2010.05.021
Zhang, M., Zhang, C. X., Pan, L. Z., Gong, S., Cui, W., Yuan, H. J., Zhang,
W. L., and Tan, J. H. (2017). Meiotic arrest with roscovitine and follicular fluid improves cytoplasmic maturation of porcine oocytes by promoting chromatin de-condensation and gene transcription. Sci. Rep. 7, 11574. doi:10.1038/S41598-017-11970-Y.