REPRODUCTIONRESEARCH

HASPIN kinase mediates histone deacetylation to regulate oocyte meiotic maturation in pigs

Zubing Cao*, Tengteng Xu*, Xu Tong*, Dandan Zhang, Chengxue Liu, Yiqing Wang, Di Gao, Lei Luo, Ling Zhang, Yunsheng Li and Yunhai Zhang

Anhui Province Key Laboratory of Local Livestock and Poultry, Genetical Resource Conservation and Breeding, College of Animal Science and Technology, Anhui Agricultural University, Hefei, China

Correspondence should be addressed to Y Zhang; Email: yunhaizhang@ahau.edu.cn

*(Z Cao, T Xu and X Tong contributed equally to this work)

Abstract

HASPIN kinase-catalyzed phosphorylation of histone H3 on threonine 3 (H3T3p) directs the activity and localization of chromosomal passenger complex (CPC) and spindle assembly checkpoint (SAC) to regulate chromosome condensation and segregation in both mitosis and meiosis. However, the function of HASPIN kinase in the meiotic maturation of porcine oocytes is not yet known. Here, we found that HASPIN mRNA is constantly expressed in porcine oocyte maturation and subsequent early embryo development. H3T3p is highly enriched on chromosomes at germinal vesicle breakdown (GVBD) stage and thereafter maintains a low level in progression through metaphase I (MI) to metaphase II (MII). Correspondingly, H3T3p was completely abolished in oocytes treated with an inhibitor of HASPIN kinase. Functionally, inhibition of HASPIN activity led to a significant reduction in the rate of oocyte meiotic maturation and the limited cumulus expansion. Additionally, HASPIN inhibition caused both spindle disorganization and chromosome misalignment in oocytes at MI and MII stage. Importantly, HASPIN inhibition severely prevented deacetylation of several highly conserved lysine (K) residues of histone H3 and H4 including H3K9, H3K14, H4K5, H4K8, H4K12 and H4K16 on the metaphase chromosomes during oocyte meiotic maturation. Taken together, these results demonstrate that HASPIN kinase regulates porcine oocyte meiotic maturation via modulating histone deacetylation.Reproduction (2019) 157 501–510

Introduction

The pig is not only an economically important domestic animal in husbandry (Whyte & Prather 2011), but also a valuable animal model for human diseases in reproductive medicine (Mordhorst & Prather 2017). Oocyte maturation (also called as meiotic maturation) in vitro is a key step of embryo production in both animal- and human-assisted reproductive technology. Nevertheless, the maturational efficiency and quality of porcine oocytes in vitro has been inferior to those derived at the physiological condition (Yuan & Krisher 2012). Thus, the better understanding of molecular mechanisms involved in controlling oocyte meiotic maturation may be beneficial to develop a novel strategy and potentially improve fertility in pigs. Numerous studies indicated that histone post-translational modifications, such as phosphorylation, acetylation, methylation, ubiquitinylation and SUMOylation, are important epigenetic factors involved in the regulation of oocyte meiotic maturation (Gu et al. 2010, Koyanagi et al. 2012, Rodriguez & Pangas 2016).

Global histone deacetylation is critical for normal chromosome condensation and segregation during

mammalian oocyte maturation (Akiyama et  al. 2006, Wang et  al. 2006, Yang et  al. 2012). The state of histone acetylation is tightly regulated by histone acetyltransferases (HATs) and histone deacetylases (HDACs). In oocyte meiotic maturation, HATs are not active, whereas HDACs dominantly execute global histone deacetylation (Kim et al. 2003, Endo et al. 2008). Previous studies indicated that inadequate histone deacetylation leads to abnormal meiotic maturation and even early embryonic lethality in several species (Akiyama et  al. 2006, Ma & Schultz 2013, Jin et  al. 2014). Although both RBBP4 and RBBP7 are reported to mediate histone deacetylation by modulating HDAC activity in mouse oocytes (Balboula et al. 2014, 2015), key epigenetic factors that regulate histone deacetylation in oocyte maturation are largely unknown. The mutual interplay between epigenetic modifications would be a promising direction as predicted by histone code. In mitosis, several studies have shown that H4K20 monomethylation and H4K16 acetylation (H4K16ac) negatively interact during mitotic cell cycle (Nishioka et  al. 2002, Rice et  al. 2002) and HASPIN-mediated H3T3p indirectly induces the deacetylation of H4K16 in yeast somatic cells (Wilkins et al. 2014). On the other

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hand, whether global histone deacetylation depends on CDC2 kinase in oocyte maturation is still controversial between mouse (Akiyama et al. 2004) and pig (Endo et al. 2006), which motivates us to continuously pursue the relationship between kinase-mediated phosphorylation and global histone deacetylation in meiosis.

Haploid germ cell-specific nuclear protein kinase (HASPIN, also called as Gsg2) was initially identified as a sperm-specific gene in mice, but its expression is also detected in cells from other tissues (Tanaka et al. 1994). HASPIN mainly catalyzes phosphorylation of histone H3 threonine 3 (H3T3p), which can be specifically blocked by a highly selective inhibitor, 5-iodotubercidin, in both somatic cells (De Antoni et al. 2012) and mouse oocytes (Nguyen et  al. 2014). Accumulating evidence has proved that HASPIN-catalyzed H3T3p is very important for multiple facets of both mitotic and meiotic progression, such as localization of chromosomal passenger complex (CPC), activation of spindle assembly checkpoint (SAC), establishment of bipolar spindle, chromosome condensation and segregation, assembly of microtubule-organizing centers (MTOCs) and cytokinesis (De Antoni et  al. 2012, Nguyen et  al. 2014, Balboula et al. 2016, Wang et al. 2016). However, the role of HASPIN kinase in the meiotic maturation of porcine oocytes is not yet known.

In the present study, we demonstrate that HASPIN and its catalyzed substrate H3T3p exist in porcine oocytes. HASPIN-mediated H3T3p is functionally required to regulate meiotic maturation, spindle organization and chromosome alignment of porcine oocytes. Importantly, HASPIN is involved in the regulation of global histone deacetylation in porcine oocyte meiotic maturation.

Materials and methods

All chemicals in this study were purchased from Sigma (Sigma-Aldrich) unless otherwise stated. All experiments were conducted in accordance with the Institutional Animal Care and Use Committee (IACUC) guidelines under current approved protocols at Anhui Agricultural University.

Preparation of HASPIN inhibitor 5-Itu

HASPIN inhibitor 5-iodotubercidin (5-Itu) (Cayman Chemical, cat no. 10010375) was dissolved in 100% ethanol. Oocyte maturation medium was used to dilute the 5-Itu stock solution to obtain the desired working solution. Because ethanol is the vehicle in which 5-Itu was dissolved, the same volume of ethanol was added into the medium as a control during oocyte maturation.

Oocyte in vitro maturation

Ovaries were collected from a local slaughterhouse and transported to the laboratory at 28–35°C in physiological saline solution. Ovaries were quickly washed in saline and antral follicles that are 3–6 mm in diameter were punctured using

a sterile 10 mL syringe with 18 gauge needles. The aspirated follicular fluid was slowly injected into a 15 mL centrifuge tube and placed on a heated stage for 15 min. Cumulus-oocyte complexes (COCs) with more than two-layer intact cumulus cells were selected under a stereomicroscope. Subsequently, COCs were cultured in one well of four-well plate containing 400 μL in vitro maturation medium (TCM-199 supplemented with 5% FBS, 10% porcine follicular fluid, 10 IU/mL eCG, 5 IU/mL hCG, 100 ng/mL l-Cysteine, 10 ng/mL EGF, 0.23 ng/mL melatonin, 2.03 × 10−5 ng/mL LIF, 2 × 10−5 ng/mL IGF-1.4 × 10−5 ng/mL FGF2, 100 U/mL penicillin and 100 mg/mL streptomycin) for 22 h, 30 h or 42 h at 38.5°C, 5% CO2 and saturated humidity. 1 mg/mL hyaluronidase in DPBS without Ca2+and Mg2+ (Gibco) was used to remove the cumulus cells surrounding oocytes at different stages. Oocytes with condensed chromosomes were considered as the occurrence of GVBD and the completion of nuclear maturation of oocytes were indicated by first polar body (pb1) extrusion.

Parthenogenetic activation of oocytes

Oocytes with pb1 extrusion were electronically activated using two pulses of direct current (1.56 kV/cm for 80 ms) in activation medium (0.3 M mannitol supplemented with 0.1 mM CaCl2, 0.1 mM MgCl2 and 0.01% polyvinyl alcohol). Subsequently, embryos were washed with PZM-3 medium three times, followed by 4 h of incubation in the chemically assisted activation medium (PZM-3 supplemented with 10 μg/mL cycloheximide and 10 μg/mL cytochalasin B). Embryos were then washed three times with PZM-3 medium and cultured in fresh PZM-3 medium at 38.5°C, 5% CO2 and 95% air with saturated humidity.

Real-time quantitative PCR

Total RNA was extracted from oocytes and embryos using RNeasy Mini Kit (Qiagen, 74104). The RNA was quantified with spectrophotometry at 260/280 nm on a NanoDrop 2000 instrument (Thermo Scientific). Reverse transcription was immediately performed using a QuantiTect Reverse Transcription Kit (Qiagen, 205311). The cDNA was aliquoted and was stored at −80°C until ready for use. The primers used in the present study are shown in Supplementary Table 1 (see section on supplementary data given at the end of this article). The assembly of PCR was prepared in FastStart SYBR Green Master (Roche, 04673514001) and was run on StepOne Plus (Applied Biosystems). The samples were collected three times and three biological replicates were conducted for each gene.

Immunofluorescence staining

Oocytes or embryos were fixed in 4% paraformaldehyde (PFA) solution for 15 min, permeabilized with 1% Triton X-100 in DPBS for 30 min at room temperature (RT) and then blocked with 2% BSA in DPBS at RT for 1 h. The samples were incubated in the blocking solution containing primary antibodies against target proteins overnight at 4°C. The detailed information regarding primary antibodies used is listed in Supplementary Table 2. After washing four times, the samples were incubated

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for 1 h in the blocking solution containing secondary antibodies in the dark at 37°C. After washing three times, the samples were counterstained for 10 min in 4,6-diamidino-2-phenylindole dihydrochloride (DAPI) or propidium iodide (PI) solution and were then loaded onto glass slides followed by being covered with a glass coverslip. Finally, the filter cubes of the inverted microscopy (Olympus) consist of narrow violet, FITC and TRITC and its camera resolution is 144 pixels per inch. The 20× objective with 1.3 of the numerical aperture (NA) was immersed in the water to image the samples at RT. The exposure power was adjusted to just below saturation relative to the group exhibiting the highest level of signal intensity and was kept constant for each oocyte in an experiment. For negative control, samples were incubated in blocking solution omitting primary antibodies.

Quantitation of histone acetylation levels

The acetylation levels of several lysine residues in oocytes were analyzed as described previously (Cao et  al. 2014). Briefly, the border around the nuclei was manually delineated according to DNA staining by ImageJ. Thereafter, at least three different cytoplasmic areas were delineated for normalization to background. The average pixel intensity of the nuclear areas was detected by ImageJ and was then normalized by dividing the average pixel intensity of the background areas to obtain the final fluorescence intensity of each acetylation modification in individual oocytes.

Statistical analysis

All experiments were carried out at least three times. The data were analyzed using one-way ANOVA or Student’s t test (SPSS 17.0) and were presented as mean ± standard error of mean (mean ± s.e.m.). P < 0.05 was considered to be statistically significant.

Results

Dynamic changes of HASPIN mRNA and H3T3p levels in porcine oocyte maturation and subsequent early embryo development

Although the function of HASPIN in both mitotic cell cycle of somatic cells (De Antoni et  al. 2012, Wang et al. 2012) and meiotic maturation of mouse oocytes (Nguyen et  al. 2014, Kang et  al. 2015, Wang et  al. 2016) has been characterized, its role in porcine oocyte meiotic maturation is still not unknown. To determine whether HASPIN mRNA is expressed in porcine oocytes and preimplantation embryos, real-time quantitative PCR was performed to examine the relative abundance of HASPIN transcripts. The results revealed that HASPIN mRNA persist during meiotic maturation and subsequent preimplantation embryo development, suggesting its maternal and zygotic origins (Fig.  1A). However, the expression levels of HASPIN are relatively lower through morula to blastocyst stage than those in both oocytes and earlier embryos (Fig.  1A) (P < 0.05). Because of the lack of porcine-specific HASPIN antibodies, we did not directly analyze the expression of HASPIN protein in oocytes. As an alternative to direct examination of HASPIN protein, immunofluorescence staining of H3T3p was performed to determine whether HASPIN kinase exists in porcine oocytes. We found that H3T3p is not detectable on the disperse chromosomes at GV, but reaches to maximal levels on the condensing chromosomes at GVBD, followed by a dramatic reduction across the aligned chromosomes upon MI and maintains low levels on the chromosomes of MII (Fig. 1B). Therefore, these data indicate that HASPIN mRNA is expressed in porcine oocyte meiotic maturation.

Figure 1 Abundance of HASPIN mRNA and H3T3p in both porcine oocytes and early embryos. (A) HASPIN expression in oocytes and early embryos. Relative abundance of HASPIN mRNA was determined by qPCR from three independent replicates. Data were normalized against endogenous housekeeping gene EF1α1 and the value from GV oocyte was set as 1. Values are shown as mean ± s.e.m. and different letters on the bars indicate significant differences (P < 0.05). (B) Levels and subcellular localization of H3T3p in oocytes at different maturational stages. Oocytes at GV, GVBD, MI and MII stages were stained for H3T3p (green) and DNA (red). Shown are representative images obtained by fluorescence microscopy. The experiment was independently repeated three times with at least 26 oocytes per stage. Bottom panel at each stage shows the merged images between H3T3p and DNA. White square insets indicate chromosomes at high magnification. White arrowhead indicates pb1. Asterisk marks chromosomes. GV, germinal vesicle; GVBD, germinal vesicle breakdown; MI,metaphase I; MII, metaphase II. Scale bar: 50 µm.

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HASPIN activity is essential for porcine oocyte meiotic maturation and cumulus expansionTo explore the function of HASPIN kinase in meiotic maturation of porcine oocytes, we matured oocytes in medium supplemented with different concentrations of 5-iodotubercidin (5-Itu), a highly selective inhibitor of HASPIN. As shown in Fig.  2A, after in vitro maturation of 44 h, the morphology of cumulus cells surrounding oocytes in the treatment groups was similar to that in control group when the dose of 5-Itu was less than and equivalent to 0.5 μM. In contrast, treatment with 1 μM 5-Itu only partially limited the cumulus expansion, but 5 μM 5-Itu completely inhibited the cumulus expansion compared to control group. Correspondingly, 5 μM 5-Itu indeed significantly reduced the rate of GVBD (Fig. 2D) and first polar body extrusion (Fig. 2B) and the area of cumulus expansion (Fig.  2C) (P < 0.05). Moreover, the developmental competence of HASPIN-inhibited oocyte with pb1 extrusion (Supplementary Fig.  1A and B) and blastocyst quality as indicated by total cell number (Supplementary Fig. 1C and D) is severely impaired. Interestingly, we note that

the optimal dose of 5-Itu required for effectively inhibiting oocyte maturation and in our subsequent experiments is analogous to that used in both somatic cells (De Antoni et  al. 2012) and mouse oocytes (Wang et  al. 2016). On the other hand, HASPIN inhibition post GVBD did not affect the rate of first polar body extrusion (Supplementary Fig.  6A), but significantly increased the rate of abnormal spindle and chromosome of oocytes at MI (Supplementary Fig.  6B) and MII (Supplementary Fig.  6C) stage and apparently reduced the developmental competence of parthenogenetic activated oocytes (Supplementary Fig.  6D). Collectively, these results suggest that HASPIN kinase is necessary for oocyte meiotic maturation and cumulus expansion in pigs.

Inhibition of HASPIN kinase effectively blocks H3T3 phosphorylation in porcine oocytes

Previous studies indicated that H3T3 is one of HASPIN substrates in other cellular contexts (Maiolica et  al. 2014). To confirm whether 5-Itu can specifically inhibit the HASPIN activity in porcine oocytes,

Figure 2 Inactivation of HASPIN kinase inhibits porcine oocyte meiotic maturation and cumulus expansion. (A) Representative images of cumulus-oocyte complexes before and after maturation. GV oocytes were matured in vitro for 44 h in the presence of the indicated concentration of iodotubercidin (5-Itu). Oocytes were matured in culture medium containing equivalent amount of ethanol and were served as a control group. (B) Comparison of the rate of oocyte maturation among different groups. The number of oocytes with first polar body after in vitro maturation for 44 h was recorded and the rate of first polar body extrusion was statistically analyzed by one-way ANOVA. The experiment was repeated six times with at least 120 oocytes per group. Values are shown as mean ± s.e.m. and different letters on the bars indicate significant differences (P < 0.05). (C) Analysis of the area of cumulus expansion among different groups. The diameter of cumulus-oocyte complexes matured in vitro for 44 h was measured and the area of cumulus expansion was calculated according to a published mathematical equation. The experiment was repeated three times with at least 30 oocytes per group. Values are shown as mean ± s.e.m. and different letters on the bars indicate significant differences (P < 0.05). (D) Comparison of the GVBD rate of oocytes. GV oocytes were randomly allocated to both control and 5-Itu-treatment group and were matured for 22 h. Cumulus cells surrounding oocytes was then mechanically removed. Denuded oocytes were stained with DAPI to observe the morphology of chromosomes by fluorescence microscopy. The number of oocytes with linear chromosomes was recorded and the GVBD rate was statistically analyzed by Student’s t test. The experiment was repeated three times with at least 90 oocytes per group. Values are shown as mean ± s.e.m. and different letters on the bars indicate significant differences (P < 0.05).

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H3T3p levels were detected by immunofluorescence staining. We observed that H3T3p on chromosomes was almost completely blocked by 5-Itu in the vast majority of oocytes at GVBD, MI and MII stage (Fig.  3A). Simultaneously, we also realized that few oocytes still contain H3T3p, whereas the proportion of oocyte with H3T3p in the treatment group is significantly reduced at each stage compared to control group (Fig.  3B, C and D) (P < 0.05). We also note that H3T3p on chromosomes in oocyte without pb1 extrusion disappear compared to control counterparts (Supplementary Fig.  5A). Therefore, our data demonstrate that HASPIN inhibition effectively abolishes H3T3 phosphorylation in porcine oocytes.

HASPIN inhibition perturbs spindle organization and chromosome alignment in porcine oocytes

To evaluate the effects of HASPIN inhibition on spindle organization and chromosome morphology, oocytes matured to MI and MII stage were stained for α-tubulin and DNA by immunofluorescence technology. We found that oocytes at MI and MII stage in control group display bipolar spindles and normal linear chromosome morphology, whereas spindles in 5-Itu-treated oocytes are disorganized, and the chromosomes are misaligned at the metaphase plate and presented as abnormal structure (Fig. 4A and B). We further observed that the proportion of MI and MII oocyte with abnormal spindles and chromosomes in the treatment group is apparently higher than that in control group (Fig.  4C, D, E and F) (P < 0.05). Therefore, these results document that HASPIN inhibition perturbs spindle organization and chromosome alignment in porcine oocytes.

HASPIN inhibition prevents deacetylation on lysine residues of histone H3 and H4 on the metaphase chromosomes of meiosis I

Given that histone deacetylation is necessary for normal oocyte meiotic maturation in several species (Ma & Schultz 2013, Jin et al. 2014, Han et al. 2015), and HASPIN-mediated H3T3p is indirectly involved in the deacetylation of H4K16 in yeast somatic cells (Wilkins et al. 2014), which prompted us to speculate that the failure to deacetylation of histone could be a main cause of abnormal meiosis in porcine oocytes. To test this hypothesis, the acetylation status of several highly conserved lysine residues of histone H3 and H4 in the metaphase chromosomes of meiosis I was analyzed by immunofluorescence staining. Firstly, the specificity of antibodies against H3K9ac, H3K14ac, H4K5ac, H4K8ac, H4K12ac and H4K16ac has been confirmed in GV oocytes (Supplementary Fig. 2B, C, D, E, F and G). To determine whether the

phenomenon of histone deacetylation exists in porcine oocyte meiotic maturation, the dynamic changes of acetylation levels on lysine residues above mentioned were characterized in detail on chromosomes of oocytes through GV to MII stage. We found that histone acetylation on all lysine residues is highly enriched in GV oocytes, and then it progressively decreases to minimal levels or even disappears in MII oocytes (Supplementary Figs 3A, B and 4A, B, C, D), suggesting that histone deacetylation indeed occur in meiotic maturation of porcine oocytes. Furthermore, acetylation levels at each lysine residue in MI oocytes treated by 5-Itu were significantly higher than those in control group (Fig. 5A, B, C, D, E and F) (P < 0.05).

Figure 3 Inhibition of HASPIN kinase effectively abolishes H3T3 phosphorylation in porcine oocytes. (A) Detection of H3T3p in oocytes at GVBD, MI and MI stage. GV oocytes were randomly distributed to control and 5-Itu treatment groups and were then matured in vitro up to a specific timing point. Oocytes at GVBD, MI and MI stage were stained for H3T3p (green) and DNA (red). Shown are representative images obtained using fluorescence microscopy. The experiment was independently repeated three times with at least 30 oocytes per group. Right panel in each group shows the merged images between H3T3p and DNA. White square insets indicate chromosomes at high magnification. GVBD, germinal vesicle breakdown; MI, metaphase I; MII, metaphase II. Scale bar: 50 µm. (B, C and D) Analysis of the proportion of oocytes with H3T3p. The number of oocytes stained positive for H3T3p between control and 5-Itu treatment groups were recorded and the percentage of oocytes with H3T3p was statistically analyzed by Student’s t test. Values are shown as mean ± s.e.m. and different letters on the bars indicate significant differences (P < 0.05).

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Altogether, these data indicate that HASPIN inhibition prevents histone deacetylation on the metaphase chromosomes of meiosis I.

HASPIN inhibition impedes deacetylation on lysine residues of histone H3 and H4 on the metaphase chromosomes of meiosis II

To further investigate whether HASPIN inhibition prevents deacetylation on lysine residues of histone H3 and H4 in the metaphase chromosomes of meiosis II, acetylation levels on each lysine residue in MII oocytes were quantitatively analyzed by immunofluorescence staining. Results revealed that acetylation levels on lysine residues were significantly elevated in 5-Itu-treated oocyte with pb1 extrusion relative to control group (Fig.  6A, B, C, D, E and F) (P < 0.05). Similarly, the increased acetylation levels were also observed in 5-Itu-treated oocyte without pb1 extrusion (Supplementary Fig.  5B). Therefore, these results indicate that HASPIN inhibition impedes histone deacetylation on the metaphase chromosomes of meiosis II.

Discussion

Despite the roles of HASPIN in both mitosis of somatic cells and meiosis of mouse oocytes being well characterized (De Antoni et  al. 2012, Nguyen et al. 2014, Wang et al. 2016), its function in porcine oocyte meiotic maturation is yet to be determined. We observe here that HASPIN is continuously expressed in porcine oocyte maturation and subsequent early embryonic development. HASPIN-catalyzed H3T3p dynamically exists on chromosomes through GVBD to MII. Additionally, HASPIN inhibition caused low maturational efficiency, spindle disorganization and chromosome misalignment of porcine oocytes. Surprisingly, HASPIN inactivation prevented global histone deacetylation in porcine oocyte meiotic maturation. Therefore, these results demonstrate that HASPIN mediates histone deacetylation to regulate the meiotic maturation of porcine oocytes.

The highly expression of porcine HASPIN transcripts persists through GV oocyte to 8-cell stage and then decreases to a low levels from morula to blastocyst stage, suggesting much more abundance of maternal Haspin mRNA in meiotic oocytes relative to mitotic embryonic cells. This differential expression pattern of HASPIN mRNA between meiotic cells and mitotic cells is similar to that observed in mouse oocytes and preimplantation embryos (Nguyen et al. 2014), indicating a conserved hallmark of HASPIN expression in oocyte maturation and subsequent embryo development among different species. Because both homologous chromosomes and sister chromatids continuously segregate in meiosis I and II while chromosome segregation occurs only once time during mitosis, it is possible that HASPIN kinase executes different functions between meiosis and mitosis.

Figure 4 HASPIN inhibition alters spindle organization and chromosome alignment in porcine oocytes. (A and B) Observation of spindle and chromosome morphology in MI and MII oocytes. GV oocytes were randomly distributed to control group and treatment group in the presence of the indicated dose of 5-Itu and were matured in vitro to MI and MII stage, respectively. Oocytes were stained for α-tubulin (green) and DNA (blue). Shown are representative images obtained using fluorescence microscopy. The experiment was independently repeated three times with at least 50 oocytes per group. Bottom panel in each group shows the merged images between α-tubulin and DNA. White square insets indicate spindles and chromosomes at high magnification. MI, metaphase I; MII, metaphase II. Scale bar: 50 µm. (C, D, E and F) Analysis of the proportion of oocytes with abnormal spindle and chromosome morphology. The morphology of spindle and chromosome in oocytes at MI and MII stage between control and 5-Itu treatment groups was scored according to a published method (Wei et al. 2014). The number of oocytes displaying abnormal spindle and chromosome morphology was recorded and the percentage of each parameter was statistically analyzed by Student’s t test. Data are shown as mean ± s.e.m. and different letters on the bars indicate significant differences (P < 0.05).

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On the other hand, we observed that H3T3p levels reach to maximum at GVBD, and then keep a low state from MI to MII. Dynamic patterns and localization of H3T3p in porcine oocyte maturation is consistent with those in mice (Wang et al. 2016). The increasing levels of H3T3p at the beginning of GVBD predict that HASPIN may be involved in the regulation of meiotic resumption. Consistent with this hypothesis is previous reported results demonstrating that HASPIN-

inhibited oocytes display the GVBD arrest (Nguyen et al. 2014, Wang et al. 2016). It is well documented that cumulus cell expansion is required for the GVBD of porcine oocytes (Yokoo et  al. 2010). The GVBD arrest of HASPIN-inhibited porcine oocytes could be attributable to defects in cumulus cell expansion and their functions. In addition, the low efficiency of first polar body of HASPIN-inhibited oocytes was also observed in this study, suggesting HASPIN-mediated

Figure 5 HASPIN inhibition causes hyperacetylation on lysine residues of histone H3 and H4 on the metaphase chromosomes of meiosis I. GV oocytes were randomly allocated to control group and treatment group in the presence of the indicated dose of 5-Itu and were matured in vitro to MI stage. Oocytes were stained for several acetylated histone lysine residues including H3K9 (A), H3K14 (B), H4K5 (C), H4K8 (D), H4K12 (E) and H4K16 (F) (red) and DNA (blue). Shown are representative images obtained using fluorescence microscopy. All experiments were repeated three times with at least 30 oocytes per group. Right panel in each group shows the merged images between histone acetylation and DNA. White square insets indicate chromosomes at high magnification. MI, metaphase I. Scale bar: 50 µm. Quantification of histone acetylation levels on the indicated lysine residues were shown to the right of each image panel. Data are shown as mean ± s.e.m. and different letters on the bars indicate significant differences (P < 0.05).

Figure 6 HASPIN inhibition leads to hyperacetylation on lysine residues of histone H3 and H4 on the metaphase chromosomes of meiosis II. GV oocytes were randomly allocated to control group and treatment group in the presence of the indicated dose of 5-Itu and were matured in vitro to MII stage. Oocytes were stained for several acetylated histone lysine residues including H3K9 (A), H3K14 (B), H4K5 (C), H4K8 (D), H4K12 (E) and H4K16 (F) (red) and DNA (blue). Shown are representative images obtained using fluorescence microscopy. All experiments were repeated three times with at least 25 oocytes per group. Right panel in each group shows the merged images between histone acetylation and DNA. MII, metaphase II. White square insets indicate chromosomes at high magnification. White arrowheads indicate pb1. Asterisks denote chromosomes. Scale bar: 50 µm. Quantification of histone acetylation levels on the indicated lysine residues were shown to the right of each image panel. Data are shown as mean ± s.e.m. and different letters on the bars indicate significant differences (P < 0.05).

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H3T3p is required for the normal progression of porcine oocyte meiotic maturation. Likewise, this phenomenon was also observed in HASPIN-inhibited mouse oocytes (Nguyen et al. 2014). Previous studies have shown that HASPIN participates in the regulation of meiotic maturation through modulating activity of CPC and SAC in mice (Nguyen et  al. 2014, Wang et al. 2016). It could thus be speculated that HASPIN inhibition leads to an inappropriate activation of CPC and SAC in porcine oocytes, in turn inducing the failure to extrude the first polar body. Meanwhile, we also found that the majority of HASPIN-inhibited porcine and mouse oocytes arrested prior to MII stage, whereas partial oocytes still could timely extrude first polar body (Nguyen et  al. 2014, Wang et  al. 2016). Unfortunately, the minor oocytes with pb1 extrusion have lost the H3T3p on chromosomes and exhibited an extremely poor developmental competence of embryo after parthenogenetic activation. This mild difference in terms of the rate of first polar body extrusion could be due to the differential responses of meiotic events in individual oocytes to HASPIN inactivation. Of note, cumulus cells play essential role for cytoplasmic maturation of oocytes (Wongsrikeao et  al. 2005, Zhang et  al. 2010). Therefore, cumulus cells are important for acquiring the developmental competence during oocyte maturation. The low developmental efficiency of embryos derived from HASPIN-inhibited oocytes could be owing to defects in cumulus cell expansion and their functions.

Many studies have demonstrated that HASPIN kinase maintains normal chromosome morphology to allow correct segregation of genetic materials in both mitosis (Higgins 2010, De Antoni et  al. 2012) and meiosis (Nguyen et al. 2014, Kang et al. 2015). In line with these findings, we found that HASPIN inhibition indeed resulted in chromosome misalignment in MI and MII oocytes. This is probably a consequence of the inability of HASPIN to drive condensin to meiotic chromosomes in oocytes (Nguyen et  al. 2014). Interestingly, HASPIN inhibition also caused spindle disorganization or even loss in MI and MII oocytes, which is consistent with that in HASPIN-inhibited mouse oocytes (Balboula et  al. 2016, Wang et  al. 2016). Recently, two studies revealed that HASPIN kinase localizes at microtubule (Wang et al. 2016) and functionally facilitates spindle assembly via regulating microtubule polymerization in mouse oocytes (Balboula et al. 2016). Therefore, these novel findings could be used to explain spindle disorganization in HAPSIN-inhibited porcine oocytes.

It is previously reported that histone deacetylation on single or several lysine residues is accompanied by oocyte meiotic maturation in multiple species, such as mouse (Kim et al. 2003), human (Huang et al. 2012), pig (Endo et al. 2005), cattle (Maalouf et al. 2008), sheep (Tang et  al. 2007), horse (Franciosi et  al.  2012) and

dog (Motheo et  al. 2017). Similarly, the deacetylation of H3K9, H3K14, H4K5, H4K8, H4K12 and H4K16 in porcine oocytes through GV to MII stage was also observed in our study. Unexpectedly, we found that HASPIN inhibition blocked the deacetylation of several lysine residues mentioned above on chromosomes in porcine MI an MII oocytes. This could be attributed to the involvement of HASPIN in the regulation of HDACs activity observed in yeast somatic cells (Wilkins et  al. 2014). As we known, histone hyperacetylation can trigger many consequences in oocyte meiotic maturation. For instance, in mice and pigs, the hyperacetylation induced by HADCs inhibitor TSA treatment, Hdac1/2 double knockout, Rbbp4/7 or Sirt6 knockdown caused the defects in some cellular events of oocyte meiotic maturation, including spindle organization, chromosome condensation and segregation, GVBD and first polar body extrusion (Akiyama et  al. 2006, Yang et al. 2012, Ma & Schultz 2013, Balboula et al. 2014, 2015, Jin et al. 2014, Han et al. 2015). In humans, insufficient deacetylation of H3K9 and H4K12 in oocyte meiotic maturation also led to aberrant spindle shapes and chromosome misalignment (van den Berg et  al. 2011, Huang et  al. 2012). Based on these studies, histone hyperacetylation induced by HASPIN inhibition could account for the abnormity of GVBD, first polar body extrusion, spindle and chromosome morphology observed in porcine oocytes. However, previous studies indicated that all classes of HDACs comprise 18 different members in cells (Zhan et  al. 2017) and are usually activated by the interaction between proteins or its phosphorylation themselves (Pflum et  al. 2001). Therefore, further investigation will be necessary to address which HDACs HASPIN kinase regulates and how HASPIN kinase modulates their activity in porcine oocyte meiotic maturation.

In conclusion, these results demonstrate that HASPIN kinase regulates oocyte meiotic maturation by promoting histone deacetylation in pigs.

Supplementary data

This is linked to the online version of the paper at https://doi.org/10.1530/REP-18-0447.

Declaration of interest

The authors declare that there is no conflict of interest that could be perceived as prejudicing the impartiality of the research reported.

Funding

This work was supported by grants from the Anhui Provincial Natural Science Foundation (1708085QC55 and

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HASPIN regulates histone deacetylation 509

https://rep.bioscientifica.com Reproduction (2019) 157 501–510

1708085MC81), The Science and Technology Major Project of Anhui province (18030701185), The Natural Science Project of Universities in Anhui Province (KJ2017A131 and KJ2016A227), The Young Scholar Natural Science Foundation of Anhui Agricultural University (2016ZR020), The High-level Talent Research Foundation of Anhui Agricultural University (YJ2016-10), The Open Fund of Anhui Province Key Laboratory of Local Livestock and Poultry, Genetical Resource Conservation and Breeding (AKLGRCB2017003), The Open Foundation of State Key Laboratory of Agrobiotechnology (2018SKLAB6-3), The Open Fund of State Key Laboratory of Genetic Resources and Evolution (GREKF18-16) and Academic & Technical Talents and the Back-up Candidates Funding for Scientific Research Activities of Anhui Province (2016H093).

Acknowledgments

The authors thank Xin Qi and Wei Ning for their help in technical assistance.

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Received 24 August 2018First decision 21 September 2018Revised manuscript received 10 March 2019Accepted 14 March 2019

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