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Reproduction (2006) 131 1037-1049
DOI: 10.1530/rep.1.00897
Copyright © 2006 Society for Reproduction and Fertility
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RESEARCH

Chromatin remodeling in somatic cells injected into mature pig oocytes

Hong-Thuy Bui1,2, Nguyen Van Thuan2, Teruhiko Wakayama2 and Takashi Miyano1

1 Department of Life Science, Graduate School of Science and Technology, Kobe University, Nada-ku, Kobe 657-8501, Japan, 2 RIKEN-Kobe, Center for Developmental Biology, Chuo-ku, Kobe 650-0047, Japan

Correspondence and reprint requests: T Miyano; Email: miyano{at}kobe-u.ac.jp


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Acknowledgements
 References
 
We examined the involvement of histone H3 modifications in the chromosome condensation and decondensation of somatic cell nuclei injected into mature pig oocytes. Nuclei of pig granulosa cells were transferred into in vitro matured intact pig oocytes, and histone H3 phosphorylation, acetylation, and methylation were examined by immunostaining with specific antibodies in relation to changes in chromosome morphology. In the condensed chromosomes of pig oocytes at metaphase II, histone H3 was phosphorylated at serine 10 (H3–S10) and serine 28 (H3–S28), and methylated at lysine 9 (H3–K9), but was not acetylated at lysine 9, 14 and 18 (H3–K9, H3–K14 and H3–K18). During the first 2 h after nuclear transfer, a series of events were observed in the somatic nuclei: nuclear membrane disassembly; chromosome condensation to form a metaphase-like configuration; an increase in histone H3 phosphorylation levels (H3–S10 and H3–S28). Next, pig oocytes injected with nuclei of somatic cells were electroactivated and the chromosome morphology of oocytes and somatic cells was examined along with histone modifications. Generally, chromosomes of the somatic cells showed a similar progression of cell cycle stage to that of oocytes, through anaphase II- and telophase II-like stages then formed pronucleus-like structures, although the morphology of the spindles differed from that of oocyte spindles. The chromosomes of somatic cells also showed changes in histone H3 dephosphorylation and reacetylation, similar to oocytes. In contrast, histone H3 methylation (H3–K9) of somatic cell nuclei did not show any significant change after injection and electroactivation of the oocytes. These results suggest that nuclear remodeling including histone H3 phosphorylation and acetylation of injected somatic nuclei took place in the oocytes under regulation by the oocyte cytoplasm.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Acknowledgements
 References
 
The successful application of mammalian cloning using differentiated adult nuclei indicates that the oocyte cytoplasm contains hitherto unidentified reprogramming activities to erase the previous memory of cell differentiation (Wilmut et al. 1997, Wakayama et al. 1998). Cloning has been achieved by introducing the nucleus of differentiated donor cells into enucleated mature oocytes. Nuclear reprogramming, which returns differentiated somatic nuclei to the totipotent undifferentiated stage, is a prerequisite for the clonal development of reconstructed embryos. The early events in this process consist of morphological remodeling of the donor nucleus, including breakdown of the nuclear membrane, initial chromatin condensation, spindle assembly, and formation of a pronucleus-like structure after activation. The timing of oocyte activation after nuclear transfer is considered to be an important factor for these remodeling events and for the efficiency of cloned embryo production (Fulka et al. 2001, Wakayama & Yanagimachi 2001). The reported time of oocyte activation varies among species: for example from immediately to 10 h after nuclear transfer in cattle (Wells et al. 1999); from immediately to 4 h in pigs (Onishi et al. 2000, Polejaeva et al. 2000), and from immediately to 6 h in mice (Wakayama et al. 1998, Fulka et al. 2001). The timing of nuclear transformation depends on the donor cell type, cell-cycle stage, and species, and the duration of direct exposure of chromosomes of somatic cells to the ooplasm may be important in reprogramming of the donor nucleus to support embryo development (Wakayama & Yanagimachi 2001). It is thought that a high level of Cdc2 kinase activity in the enucleated mature oocyte induces breakdown of the nuclear membrane of the transferred nucleus, resulting in chromosome condensation (Campbell et al. 1996, Tani et al. 2001). When a stimulus for activation is applied, the activity of Cdc2 kinase decreases, resulting in chromosome decondensation and formation of a pronucleus-like structure. During the activation of intact mature oocytes, each pair of sister chromatids with two kinetochores separates: one set remains in the oocyte, and the other is extruded into the second polar body. However, the kinetics of the condensed chromosomes derived from injected somatic nuclei is not well understood: each of the somatic chromosomes has only one kinetochore because the nuclei are derived from cells in the G0/G1 phase of the cell cycle.

The DNA is packaged with histone proteins into chromatin, compacting DNA some 10 000-fold compared with interphase. Four core histone molecules, H2A, H2B, H3, and H4, associate with DNA to form the basic nucleosome structure in which DNA wraps around the histone proteins in eukaryotic cells (Arents et al. 1991). It has been suggested that histones are major carriers of epigenetic information, and that covalent modifications on the histone N-terminal tails function as master on/off switches that determine whether a gene is active or inactive (Fischle et al. 2003). Histone tails are subjected to a wide range of posttranslational modifications, including acetylation, phosphorylation, and methylation (Strahl & Allis 2000). These modifications of histone H3 are thought to play important roles in the regulation of gene expression in reconstructed embryos and to be essential for their development. Thus, histone H3 is acetylated at lysine (K) 9, 14, 18 and 23, phosphorylated at serine (S) 10 and 28, and methylated at K4 and K9 (Cheung et al. 2000). The juxtaposition of these sites provides potential cross-regulation of different modification events. In our previous experiments, histone H3 was shown to be phosphorylated at S10 in condensed chromosomes during the maturation of pig oocytes, and chromosome condensation was associated with histone H3 kinase activity (Bui et al. 2004). In somatic cells, it has been suggested that the phosphorylation of histone H3 at S10 may be linked with the acetylation of histone H3 at K14 during prophase (Mateescu et al. 2004). In addition, histone H3 phosphorylation at S28 occurs independently of phosphorylation at S10 following stimulation of the Ras-MAPK (mitogen-activated protein kinase) pathway (Dunn & Davie 2005). Although Cdc2 kinase and MAPK are active in mature oocytes, it is not known whether histone H3–S28 phosphorylation occurs in the oocytes. Moreover, histone H3 in oocytes is uniquely acetylated. For example, the acetylation of histone H3–K9 persists during the mitotic phase in somatic cells (Kruhlak et al. 2001), although the acetylation of histone H3–K9 and H3–K14 is removed during the meiotic maturation of mouse oocytes (Kim et al. 2003). Furthermore, histone H3-K9 is trimethylated in mouse oocytes, which has been a putative epigenetic marker (Arney et al. 2002, Liu et al. 2004). After nuclear transfer of somatic cells at G0/G1 phase into the oocytes at metaphase II, somatic nuclei form condensed chromosomes, but it remains to be elucidated how their histone modifications change in the somatic nuclei under the influence of the ooplasm. In addition, after oocyte activation, oocyte chromosomes decondense to form chromatin. It is not yet known how histone modifications of the somatic nuclei change in activated oocytes. It is possible that such changes in histone modifications of the somatic nuclei affect the efficiency of cloning by somatic cell nuclear transfer.

Here we aimed to determine the involvement of histone H3 phosphorylation, acetylation, and methylation in the morphological changes of somatic nucleus chromosomes following transfer into in vitro matured intact pig oocytes. Injected pig oocytes were electro-activated and the chromosome morphology of both the oocytes and injected somatic cell nuclei were examined in relation to histone modifications in the same ooplasm.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Acknowledgements
 References
 
Collection and maturation culture of oocytes
Pig ovaries were obtained from prepubertal gilts at a local slaughterhouse. The ovaries were washed once with 0.2% (w/v) cetyltrimethylammonium bromide and twice with Dulbecco’s phosphate-buffered saline (PBS) containing 0.1% (w/v) polyvinyl alcohol (PBS–PVA Sigma-Aldrich, St. Louis, MO, USA). Antral follicles 4–6 mm in diameter were dissected in PBS–PVA from ovaries following the technique described by Moor and Trounson (1977). After opening the follicles in 25 mM HEPES buffered TCM-199 (Earl’s salts; Nissui Pharmaceutical, Tokyo, Japan), oocyte–cumulus complexes with a piece of parietal granulosa tissue (oocyte–cumulus–granulosa cell complexes; OCGCs) were isolated from the follicles. Following two washes in HEPES-buffered TCM-199, the OCGCs were cultured for 45 h in 2 ml of bicarbonate-buffered TCM-199 supplemented with 10% (v/v) heat-treated fetal calf serum (Biocell Inc., Carson, CA, USA), 0.1 mg/ml sodium pyruvate, 0.08 mg/ml kanamycin sulfate (Sigma-Aldrich), 2.2 mg/ml sodium bicarbonate, 0.1 IU/ml human menopausal gonadotropin (Pergonal; Teikoku Zoki, Tokyo, Japan), and two everted theca shells, using gentle agitation in an atmosphere of 5% CO2 in humidified air at 38.5 °C. After culture, oocytes were completely denuded from cumulus cells using 0.1% hyaluronidase (Sigma-Aldrich) and gentle pipetting, and were then used for nuclear transfer experiments.

Collection and culture of somatic cells
Healthy pig antral follicles 4–6 mm in diameter were everted in HEPES-199 using two pairs of fine forceps to collect mural granulosa cells. The cells were dispersed by vigorously aspirating them in and out of a pipette, and the suspension was then transferred into a 1.5 ml tube. After centrifugation at 500 g for 3 min, the pellet was resuspended in PBS. This treatment was repeated twice. The resulting pellet of granulosa cells was resuspended in a small amount of Dulbecco’s modified Eagle’s medium (DMEM, Sigma-Aldrich) supplemented with 10% fetal calf serum, and the cells were seeded in a dish whose bottom had been covered by 0.1% gelatin (Sigma-Aldrich). Next, 2 ml of DMEM were added to the culture dish. The cells were cultured in an atmosphere of 5% CO2 in humidified air at 38.5 °C for 2 days. After culture, cells were collected by trypsinization. Briefly, the culture medium was removed and the cells were thoroughly rinsed twice with PBS. Trypsin solution (Sigma-Aldrich) was added to cover the cultured granulosa cells for 1 min, and then 2 ml of serum-supplemented DMEM were added. The dispersed granulosa cells were transferred into a tube. After three washes and centrifugation in PBS, they were suspended in HEPES–CZB medium (Chatot et al. 1989).

Nuclear transfer and oocyte activation
Nuclear transfer of granulosa cells was carried out using a piezoactuated micromanipulator system (Primetech Corp., Tsuchiura, Japan). Oocytes with the first polar body at metaphase II (MII) were selected and placed in 10 µl droplets of HEPES–CZB in a micromanipulation chamber. Trypsinized granulosa cells were transferred into droplets of HEPES–CZB containing 12% polyvinyl pyrrolidone (PVP, Mr 360 kDa; Wako, Japan). Each of the oocytes was stabilized using a holding pipette at the 9 o’clock position and then rotated until the first polar body was found either at the 12 or the 6 o’clock position. Because the MII spindle is generally close to the first polar body, such positioning was aimed to avoid damage to the MII spindle. The plasma membrane of the granulosa cell was removed using an injection pipette (6–8 µm inner diameter) with several piezo pulses, and a single granulosa cell nucleus was injected from the 3 o’clock position. After nuclear injection, oocytes were cultured in bicarbonate-buffered TCM-199 supplemented with 10% fetal calf serum, 0.1 mg/ml sodium pyruvate, 0.08 mg/ml kanamycin sulfate, and 2.2 mg/ml sodium bicarbonate. After culturing for 0.5, 1, 2, 3, 4, and 5 h, oocytes were fixed for immunofluorescence microscopy (below).

Another group of injected oocytes was cultured in the same medium for 2 h before electroactivation. Oocytes were activated using a protocol described previously (Nguyen et al. 2003). They were washed three times in a solution comprising 0.3 mM mannitol, 0.1 mM MgSO4, 0.05 mM CaCl2 and 0.01% (w/v) PVA. Each group (< 20 oocytes) was transferred to 100 µl of the solution between two parallel stainless electrodes in a chamber (FTC-03; Shimadzu Co. Ltd., Kyoto, Japan). A single direct-current pulse of 1 500 V/cm for 100 µs was supplied from an Electro Cell Manipulator (ECM 2000; BTX Inc., San Diego, CA, USA). Electroactivated oocytes were then washed three times in HEPES-199 to rinse off the mannitol solution and cultured for 2, 4, and 6 h in bicarbonate-buffered TCM-199 supplemented with 10% fetal calf serum, 0.1 mg/ml sodium pyruvate, 0.08 mg/ml kanamycin sulfate, and 2.2 mg/ml sodium bicarbonate. The activated oocytes were cultured under the same conditions used for oocyte maturation. After culturing, the oocytes were fixed and used for immunofluorescence microscopy (below).

Immunofluorescence microscopy
Oocytes collected after culture were washed twice in PBS–PVA and fixed in PBS–PVA containing 4% (w/v) paraformaldehyde and 0.2% (v/v) Triton X-100 for 40 min. The fixed oocytes were washed twice in PBS–PVA for 15 min each and stored overnight in PBS–PVA with 1% (w/v) bovine serum albumin (International Regents Corporation, Kobe, Japan; BSA–PBS–PVA) at 4 °C. The oocytes were blocked with 10% (v/v) goat serum (DakoCytomation A/S, Glostrup, Denmark) in BSA–PBS–PVA for 45 min and then incubated with the first antibodies at 4 °C overnight. The primary antibodies used here were rabbit polyclonal anti-phospho-histone H3 at serine position 10 (Cell Signaling Technology Inc., MA, USA), rabbit polyclonal anti-phospho-histone H3 at serine 28 (Upstate Cell Signaling Solutions, VA, USA), rabbit polyclonal anti-trimethyl-histone H3 at lysine 9 (Abcam, Cambridge, UK), rabbit polyclonal anti-acetyl-histone H3 at lysine 9 (Upstate), rabbit polyclonal anti-acetyl-histone H3 at lysine 14 (Upstate), and rabbit polyclonal anti-acetyl-histone H3 at lysine 18 (Abcam) antibodies. To determine the integrity of the nuclear membrane and meiotic stages, mouse monoclonal anti-lamin A/C (Santa Cruz Biotechnology Inc., CA, USA) and mouse monoclonal anti-{alpha} tubulin (Sigma-Aldrich) antibodies were used. After being washed three times in BSA–PBS–PVA for 15 min each, the oocytes were incubated with secondary antibodies: Alexa Fluor 488-labeled goat anti-rabbit IgG (Molecular Probes Inc., Eugene, OR, USA) and Alexa Fluor 568-labeled goat anti-mouse IgG (Molecular Probes) for 40 min at room temperature. After being washed three times in BSA–PBS–PVA for 15 min each, the chromosomes were stained with 4,6-diamidino-2-phenylindole (DAPI) (2 µg/ml; Molecular Probes). Following complete washing, the oocytes were mounted on slides using Vectashield mounting medium (Vector Laboratories Inc., Burlingame, CA, USA). Observations were carried out using a confocal scanning laser microscope (Radiance 2100, Bio-Rad, Hercules, CA, USA). The somatic cell nucleus was distinguished from the oocyte nucleus according to the positions of the polar bodies. For negative controls, oocytes were reacted with nonimmune rabbit serum instead of rabbit polyclonal first antibodies, or with mouse IgG instead of mouse monoclonal antibodies.

Statistical analysis
More than 30 immunostained oocytes were examined in each group. The numbers of oocytes were analyzed using one-way ANOVA followed by Tukey’s test for multiple comparisons; P < 0.05 was considered statistically significant. For quantitative analysis, the fluorescence images were subjected to densitometric analysis using Image-J software from the National Institutes of Health (http://rsb.info.nih.gov/ij/).


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Acknowledgements
 References
 
Chromosome morphology and histone H3 modifications of somatic cell nuclei after injection into mature oocytes
Pig oocytes maintained their MII stage for 5 h after nuclear transfer of granulosa cells (Tables 1Go and 2Go, and Fig. 1Go). Histone H3 phosphorylation at serine 10 (S10) and serine 28 (S28) in the oocyte chromosomes was also maintained during the culture period.


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  		Table 1 Chromatin morphology and phosphorylation of histone H3–S10 in somatic cell nuclei transferred into MII pig oocytes.
 

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  		Table 2 Chromatin morphology and phosphorylation of histone H3–S28 in somatic cell nuclei transferred into MII pig oocytes.
 

Figure 1
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  		Figure 1 Changes in the histone phosphorylation of pig granulosa cell nuclei transferred into MII pig oocytes. Nuclei of pig granulosa cells were injected into MII oocytes. Oocytes were examined after 0–5 h by immunostaining with Alexa 488-labeled anti-phospho-histone H3–S28 (P-H3–S28) antibody (green). Microtubules were stained with Alexa 568-labeled anti-{alpha} tubulin antibody (red). The DNA was counterstained with DAPI (blue). Key: ON, oocyte nucleus; SN, somatic cell nucleus; 1st pb, first polar body. Figures B1 and C1 show the somatic cell nuclei at high magnification, and B2 shows the oocyte nucleus. Scale bar =10 µm.

 
Phosphorylations of histones H3–S10 and H3–S28 were not detected in any somatic cell nuclei immediately after injection (Fig. 1Go.A' for P-H3–S28). Chromatin of injected somatic cell nuclei began to condense to form a single cluster of chromosomes, as in the diakinesis stage of oocyte maturation, 0.5 h after nuclear transfer (Tables 1Go and 2Go). In the cluster of condensed chromosomes, histone H3 began to be phosphorylated at both positions of S10 and S28 in 49% and 55% of oocytes respectively. Some somatic cell nuclei appeared to be in a metaphase-like stage, but there were only a few at 1 h. At this time, the intensity of the fluorescent signal of phosphorylated histone H3 was low at both positions S10 and S28. After 2 h, the proportion of somatic cell nuclei showing metaphase-like chromosomes had increased to approximately 80% (Fig. 1Go); thereafter, a high proportion was maintained until 5 h after nuclear transfer. High levels of phosphorylation of H3–S10 and H3–S28 in the condensed chromosomes of somatic cells were observed in more than 90% of oocytes between 2 and 5 h after nuclear transfer.

There were some differences between the morphologies of metaphase chromosomes of oocyte and somatic nuclei. The oocyte chromosomes were located in the metaphase plate at the equator of the spindle (Figs. 1.B and 1.BGo2), whereas somatic chromosomes were located on spindle poles or were dispersed on the spindle (Fig. 1.BGo1). In addition, in some oocytes the chromosomes of somatic cells were dispersed in the ooplasm, away from the spindle (Fig. 1.CGo1), while some others had a cluster of somatic cell chromosomes even 5 h after nuclear transfer. Histones H3–S10 and H3–S28 were phosphorylated in the clumps of condensed chromosomes (Tables 1Go and 2Go).

Fluorescent signals of histone H3 acetylation (Ac-H3) were detected for lysine 9 (K9), lysine 14 (K14), and lysine 18 (K18) in the somatic cell nuclei immediately after nuclear transfer, whereas no acetylation signal was observed in oocyte chromosomes (Table 3Go). The intensity of the signal of Ac-H3–K9 in somatic cell nuclei decreased to a low level at 0.5 h and disappeared 1 h after nuclear transfer. The signals of Ac-H3–K14 and Ac-H3–K18 decreased after 1 h and disappeared after 2 h (Table 3Go).


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  		Table 3 Acetylation of histone H3 at K9, K14 and K18, and methylation of histone H3 at K9 in somatic cell nuclei transferred into MII pig oocytes.
 
The integrity of the nuclear membranes of injected somatic cell nuclei was monitored by immunostaining with anti-lamin A/C antibody. After 0.5 h, the antibody recognized the entire contour of the somatic cell nuclei (Table 3Go). After 1 h, a breakdown of the nuclear membrane was observed in some somatic cell nuclei. After 2 h, most of the nuclear membranes were completely disassembled.

The signal from the antibody that recognized the trimethylated form of histone H3 at lysine 9 (Me-H3–K9) was detected in both oocyte and somatic cell nuclei immediately after injection. This was maintained for 5 h after nuclear transfer (Table 3Go).

The negative control oocytes did not show any fluorescent signals at any stage.

Chromosome morphology and histone H3 modifications of injected somatic cell nuclei after electroactivation of oocytes
The proportion of somatic cell nuclei showing meta-phase-like chromosomes reached a peak 2 h after nuclear transfer, and there was no statistical difference in the proportions at 2 h or later (Tables 1Go and 2Go). Therefore, injected oocytes were electroactivated 2 h after nuclear transfer to examine the changes in chromosome morphology and their histone modifications after oocyte activation.

Two hours after activation, approximately 90% of oocytes were at anaphase II/telophase II (AII-TII), and chromosomes of the somatic cell nuclei in 50% of oocytes showed anaphase/telophase (A-T)-like morphology (Figs. 2Go and 3Go). The proportion increased to approximately 70% at 4 h after activation. At this time, decondensation of the chromosomes started in some oocytes. After 6 h, oocyte chromosomes completely decondensed to form a female pronucleus, and somatic chromosomes also formed a pronucleus-like structure.


Figure 2
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  		Figure 2 Chromosome morphology and histone H3 dephosphorylation of oocyte and injected somatic cell nuclei after electroactivation. Nuclei of pig granulosa cells were injected into MII oocytes. Microinjected oocytes were cultured for 2 h, electroactivated and examined after 0, 2, 4, and 6 h by immunostaining with Alexa 488-labeled specific antibodies to phospho-histone H3–S10 (Fig. 2.I, P-H3–S10) and phospho-histone H3–S28 (Fig. 2.II, P-H3–S28). The DNA was counterstained with DAPI. The columns and bars represent means±S.E.M.S of five replicate experiments. The phosphorylation level of histones in oocyte chromosomes at MII was used as a control and set at 100%. The fluorescence intensity observed for each sample was expressed relative to this value. Intensity > 60% indicates highly phosphorylated histone H3 (+); intensity 20–60%, moderately phosphorylated histone H3 (+/–); intensity < 20%, dephosphorylated histone H3.

 

Figure 3
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  		Figure 3 Dephosphorylation of histone H3–S10 (Fig. 3.I) and H3–S28 (Fig. 3.II), of somatic cell chromosomes in electroactivated MII pig oocytes. Microinjected oocytes were cultured for 2 h and then electroactivated. Oocytes were examined after 0 h (2 h after injection of somatic nuclei as shown in Fig. 1.B and 1.C), 2, 4, and 6 h by immunostaining with specific Alexa 488-labeled antibodies to phospho-histone H3–S10 and phospho-histone H3–S28 (green). The DNA was counterstained with DAPI (blue). Key: SN, somatic cell nucleus; ON, oocyte nucleus; 1st pb, first polar body; SPN, somatic cell pronucleus, and FPN, female pronucleus. Meiotic stages of oocytes in A, B, and C are anaphase II, telophase II, and the pronucleus stage respectively. Scale bar=20 µm.

 
The phosphorylation levels of H3–S10 and H3–S28 began to decrease in both oocyte and somatic chromosomes 2 h after electroactivation (Fig. 2Go). The phosphorylation levels of somatic chromosomes decreased earlier than those of oocyte chromosomes but this difference was not statistically significant. The phosphorylation of H3–S10 disappeared completely when the chromosomes had decondensed in both the female pronucleus and somatic pronucleus 6 h after electro-activation (Fig. 3.I.C'Go). However, the phosphorylation of histone H3–S28 was still maintained at low levels at the periphery of the nucleoli in both pronuclei (Fig. 3.II.C'Go), and had completely disappeared at the 2-cell stage (data not shown).

Before pronuclear formation, the spindles of somatic cells immunostained with the anti-{alpha} tubulin antibody showed different morphologies from those of oocytes (Fig. 4Go). Somatic chromosomes in the spindles showed three arrangements: some had one cluster (17%, Figs. 4.I.AGo1 and 4.II), some had two clusters (54%, Fig. 4-I.CGo1), and some had more than two clusters (29%, Fig. 4-I.BGo1), whereas all of the oocyte chromosomes formed two clusters at the MII–pronucleus stage transition (ON in Figs. 4.I.A, B, and CGo). The formation of pronuclei also differed between somatic and oocyte chromosomes. In some oocytes, somatic chromosomes formed 2 or more pronuclei (Figs. 4.I.DGo1 and 4.II), while oocytes formed only one female pronucleus (FPN in Fig. 4.I.DGo). Generally, the somatic chromosomes showed a similar cell-cycle stage to that of oocytes. In some cases, the oocyte chromosomes developed further, whereas somatic chromosomes remained clumped (Figs. 4.I.A and 4.I.AGo1). However, the phosphorylation and dephosphorylation patterns of histones H3–S10 and H3–S28 in the somatic chromosomes changed in a manner similar to that in oocyte chromosomes.


Figure 4
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  		Figure 4 Somatic cell chromosomes and pronuclei showing different morphologies from those of pig oocytes. Nuclei of pig granulosa cells were injected into MII oocytes. Subsequently, the oocytes were cultured for 2 h and then electroactivated. Oocytes were examined after 2, 4, and 6 h by immunostaining with Alexa 488-labeled anti-phospho-histone H3–S10 (P-H3–S10) antibody (green), and Alexa 568-labeled anti-{alpha} tubulin antibody for the microtubules (red). The DNA was counterstained with DAPI (blue) in Fig. 4.I. Somatic cell chromosomes are shown in Figs. 4.A1–4.D1 at high magnification (merged images). The distribution of different types of morphologies of somatic cell chromosomes and pronuclei between 2 h and 6 h after electroactivation is shown in Fig. 4.II. Oocytes were divided into two classes, before (n=72) and after (n=69) pronuclear formation. The morphologies of somatic cell and oocyte nuclei were categorized by the numbers of chromosome clusters or pronuclei. Key: SN, somatic cell nucleus; ON, oocyte nucleus; 1st pb, first polar body; SPN, somatic pronucleus, and FPN, female pronucleus. Scale bar =20 µm.

 
There were no fluorescent signals of any acetylated lysine in oocyte chromosomes or injected somatic nuclei 2 h after nuclear transfer (0 h of activation; Table 3Go). Acetylation of H3–K9 and H3–K14 reappeared in oocyte chromosomes at the AII-TII stage (Table 4Go). Thereafter, acetylation of H3–K18 appeared in oocytes at the pronucleus stage (Fig. 5Go). The acetylation of H3 at K9, K14, and K18 in somatic chromosomes showed similar changes to those in oocyte chromosomes. However, in 10–20% of somatic chromosomes there was no reappearance of H3 acetylation, even though the oocyte chromosomes were reacetylated.


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  		Table 4 Reacetylation of histone H3 at K9, K14 and K18, and methylation of histones H3 at K9 in somatic cell nuclei transferred into MII pig oocytes.
 

Figure 5
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  		Figure 5 Chromosome morphology and histone H3 reacetylation of oocyte and injected somatic cell nuclei after electroactivation. Nuclei of pig granulosa cells were injected into MII oocytes. Subsequently, the oocytes were cultured for 2 h and then electroactivated. Oocytes were examined after 2, 4, and 6 h by immunostaining with Alexa 488-labeled anti-acetyl-histone H3–K18 (Ac-H3–K18) antibody (green), and Alexa 568-labeled anti-{alpha} tubulin antibody for the microtubules (red). The DNA was counterstained with DAPI (blue). Key: 1st pb, first polar body; 2nd pb, second polar body; SN, somatic cell nucleus; ON, oocyte nucleus; SPN, somatic pronucleus; FPN, female pronucleus. Scale bar=20 µm.

 
After electroactivation, methylation of H3–K9 was maintained in both oocyte and somatic chromosomes until the pronucleus stage (Table 4Go).


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Acknowledgements
 References
 
Arrest of pig oocytes at the MII stage was maintained for 5 h after nuclear transfer of somatic cells. During this time, remodeling of pig granulosa cell nuclei took place in the oocytes. A series of events were observed during the first 2 h, including nuclear membrane disassembly, chromosome condensation to form a metaphase-like configuration, a decrease in histone H3 acetylation levels, and an increase in histone H3 phosphorylation levels. These results indicate that the nuclear stage and modifications of histone H3 in injected somatic nuclei are controlled by the cytoplasm of MII oocytes.

We have reported that oocytes at the MII stage have high activity of both Cdc2 and histone H3 kinases, and have suggested that activation of Cdc2 kinase and histone H3 kinase is required for the nuclear membrane breakdown and chromosome condensation of pig oocytes respectively (Bui et al. 2004). It is thought that the nuclear membrane of injected somatic cells is broken down by Cdc2 kinase, and that histone H3 phosphorylation and chromosome condensation are induced by histone H3 kinase activity and MAPK which is also activated in MII oocytes and has the ability to phosphorylate histone H3 (Bui et al. 2004), although the relation among these kinases has not been elucidated. In 80% of oocytes, somatic cell nuclei formed metaphase-like spindles 2 h after nuclear transfer. However, the shape was different from that of oocytes. Oocyte chromosomes were aligned at the spindle equator, whereas the chromosomes of somatic cells were located on spindle poles or on the spindle, as previously shown in the mouse (Wakayama et al. 1998). They sometimes dispersed in the ooplasm away from the spindle. It has been suggested that chromatin has the ability to stabilize microtubules and to organize them into bipolar spindles independently of kinetochores and centrosomes (Dogterom et al. 1996, Heald et al. 1996). Thus, magnetic beads coated with plasmid DNA have been shown to induce bipolar spindle assembly in Xenopus egg M-phase extracts (Heald et al. 1996). Pig oocytes have no centriole and form centrosome-free meiotic spindles (Lee et al. 2000).

However, the somatic cell at G0/G1 has a centrosome (a pair of centrioles) located near the nucleus. The centrosome is the microtubule-organizing center, and forms the spindle pole during mitotic division of somatic cells. In the present experiment, if the somatic nuclei contained a centrosome at the periphery of the nuclear membrane, it would happen the centrosome was introduced into the oocytes. It is possible that the centrosome caused the abnormal spindle formation in centrosome-free pig oocytes. Alternatively, the chromosomes derived from injected granulosa cells at the G0/G1 phase were monovalents, which are thought to be unable to establish correct alignment in the spindle. Oocyte chromosomes at the MII stage consist of two sister chromatids, each of which goes to the opposite spindle pole when the chromosomes are separated at the onset of AII. The abnormalities in the somatic spindles would cause unequal separation of somatic chromosomes after electroactivation of the oocytes.

Although the spindle morphology of somatic cell nuclei was found to be different from oocyte nuclei, they showed anaphase- and telophase-like stages similar to those of oocytes after electroactivation. However, during these stages, somatic nuclei formed two or more clusters of chromosomes. This abnormality would result in some oocytes having more than two somatic pronuclei, for example one large and two small pronuclei after electroactivation, as described previously for the mouse (Kishikawa et al. 1999). On the other hand, the dephosphorylation of histone H3–S10 and H3–S28 of the somatic chromosomes took place in a similar manner to oocyte chromosomes in the pronucleus-like structures, which suggests that dephosphorylation of histone H3 in injected somatic nuclei is controlled by the oocyte cytoplasm.

It has been thought that the cytoplasm of MII oocytes is able to reprogram the gene expression pattern of transferred somatic cell nuclei (Wilmut et al. 1997, Kikyo & Wolffe 2000). Histone acetylation in chromatin is thought to be associated with increased gene expression (Tse et al. 1998). Schultz et al.(1999) have suggested that the oocyte genome must be reprogrammed during meiosis to allow the remarkable transformation from differentiated oocytes into the totipotent embryos of the next generation. In the present experiments, no examined histone H3 acetylation was detected in the chromosomes of MII oocytes; perhaps this was caused by cytoplasmic activity to deacetylate the histones (Kim et al. 2003). Deacetylation of histones in oocytes may be involved in genome remodeling by erasing the epigenetic markers for active genes in the oocytes (Kim et al. 2003). We found that histone H3 molecules in somatic cell nuclei were acetylated at all the examined positions immediately after transfer. However, the acetylation levels decreased and disappeared completely 2 h after nuclear transfer. Thus, the cytoplasm of MII oocytes could remove the acetyl groups from H3–K9, H3–K14, and H3–K18 in the transferred somatic cell nuclei. It is likely that the deacetylation is involved in erasing the epigenetic memory of differentiated somatic cells.

Transcription in pig oocytes is initiated at the 4-cell stage (the third cell cycle after fertilization), when functional nucleoli develop (Maddox-Hyttel et al. 2001). In addition to the regulation of transcription, histone acetylation has been thought to be associated with DNA replication (Vogelauer et al. 2002). After electroactivation, acetylation of H3–K9/H3–K14 and H3–K18 of oocyte chromosomes reappeared at AII–TII and at the pronucleus stage respectively perhaps in preparation for later DNA replication and future transcriptional activity. During the reacetylation of H3–K9 and H3–K14 at AII–TII, the phosphorylation levels of histone H3–S10 decreased rapidly and in some oocytes H3–S10 disappeared in this stage. This finding suggests that the reacetylation of H3–K9 and H3–K14, and dephosphorylation of H3–S10 possibly interact in the oocytes. This hypothesis is supported by the report based on budding yeasts and in nematodes where a decrease in the acetylation of H3–K9 was observed when H3–S10 phosphorylation levels increased (Hsu et al. 2000). However, in somatic cells, mitogenic stimulation induces rapid phosphorylation of histone H3–S10 and synergistic acetylation of histone H3 (Cheung et al. 2000). Furthermore, there is a positive correlation between phosphorylation of histone H3–S10 and acetylation of histone H3–K14, which promotes delocalization of heterochromatin protein 1 (HP1) from the chromatin at the G2 phase (Mateescu et al. 2004). This HP1 delocalization from chromatin has been observed at the G2/M transition of somatic cells (Murzina et al. 1999). Granulosa cell nuclei used in the present study underwent rapid chromosome condensation after injection into MII oocytes. The chronology in both oocyte and injected somatic cell nuclei that we observed suggests that the phosphorylation of histone H3–S10 occurred corresponding with histone H3 deacetylation (H3–K9 and H3–K14), and that after electroactivation histone H3 reacetylation occurred in concert with H3–S10 dephosphorylation. This means that the somatic histone modifications in the oocytes are different from those in normal somatic cell cycle, and that the phosphorylation and acetylation of histone H3 in injected somatic cell nuclei are completely controlled by the ooplasm, and not by the somatic nucleus itself. This MII-oocyte specific ability of histone deacetylation is perhaps attributed to the meiotic specific histone deacetylase (HDAC) activity, which decreases soon after oocyte activation (Kim et al. 2003).

In the pronuclear stage, histone H3 was completely dephosphorylated at S10 but was still phosphorylated at S28 around the nucleoli. Finally, phosphorylation of H3–K28 completely disappeared at the 2-cell stage when histone H3 became intensely acetylated (data not shown). The result suggests that phosphorylational events of H3–S10 and H3–S28 are regulated separately in activated pig oocytes. It has been shown in immunolocalization studies of mouse somatic cells that phosphorylation events of histones H3–S10 and H3–S28 occur independently in distinct chromatin regions, and that H3–S28 phosphorylation has a higher steady state of H3–K14 acetylation than that of H3–S10 (Dunn & Davie 2005). After electroactivation of pig oocytes, rapid reacetylation occurred at all examined lysine residues. Persistence of phosphorylation of H3–S28 might be due to the interaction with acetylated H3–K14 in the oocytes that started the somatic cell cycle.

In normal mammalian fertilization, the oocyte cytoplasm regulates and enhances the epigenetic asymmetry between parental genomes so that functional differences are observed between genomes during development. Epigenetic differences are enhanced in the zygote by means of DNA demethylation of the paternal genome shortly after fertilization in mice (Mayer et al. 2000), at the pronucleus stage in pigs (Gioia et al. 2005), or within the first embryonic cell cycle in sheep (Young & Beaujean 2004), while the maternal genome displays de novo methylation. Santos et al.(2003) suggested that histone H3–K9 methylation may be reprogrammed in parallel with DNA methylation in bovine embryos. Liu et al.(2004) suggested that histone H3 methylase is active in oocytes before fertilization, but not afterwards, and that this asymmetric methylation pattern is generated by the change in methylase activity in the cytoplasm after fertilization in mice. Therefore, mouse zygotes show a high level of histone H3–K9 methylation in the maternal chromosomes, whereas the paternal chromosome has no such methylation at this position. In the present experiments, methylation of H3–K9 was observed in the somatic cell nuclei; this did not change by 2 h after nuclear transfer and was not diminished after subsequent activation in pig oocytes. From this result we think that, if embryos are created by somatic cell nuclear transfer, they should have a set of chromosomes with highly and differently methylated histones compared with normal fertilization. This persistence of histone methylation in transferred somatic nuclei in our study is good agreement with the observation that bovine nuclear transfer embryos show hypermethylation of histone H3–K9 associated with DNA methylation (Santos et al. 2003).

The methylation of H3 at K9 persists through the time course after nuclear transfer and electroactivation regardless of the deacetylation and reacetylation of H3–K9 and the phosphorylation and dephosphorylation of H3–S10 in the somatic cell nuclei. Histone H3–K9 of MII pig oocytes was methylated and H3–S10 was phosphorylated, and histone H3 of injected somatic nucleus showed similar modificational change. This K9/S10 modification is a putative ‘methyl/phospho switch’ serving to diminish the binding affinity of HP1 to methylated H3–K9 (Fischle et al. 2003). This modification possibly concerned removal of HP1 and rapid condensation of somatic chromosomes in the oocytes. At fertilization, sperm-specific protamines are replaced by oocyte-cytoplasmic histones (McLay & Clarke 2003). The hypo-methylation of histone H3–K9 in male pronucleus as described above implies that oocyte-cytoplasmic histone H3 is demethylated at K9 and/or the histone methylase activity in the oocytes is low in fertilized oocytes. It was reported recently that the unmethylated male pronucleus underwent de novo methylation when it was transferred into enucleated GV- or MII-stage mouse oocytes, which suggests that histone H3 methylase is active before fertilization, but not afterwards, and that the asymmetric methylation pattern is generated by this change in methylase activity in the cytoplasm after fertilization (Liu et al. 2004). In the present study, methylation of histone H3–K9 of oocyte and somatic cell nuclei was maintained after oocyte activation. We speculate that the methylated histone H3 molecules before activation are not or rarely exchanged with cytoplasmic unmethylated histone H3 molecules in the activated oocytes.

In summary, we compared the morphology between oocyte and somatic cell chromosomes with histone modifications in the same cytoplasm of mature oocytes. The somatic cell chromosomes showed similar morphological changes to those of oocytes. Although the spindle morphology was different between oocyte and somatic cells, histone H3–S10 and H3–S28 in the somatic cell chromosomes were phosphorylated and dephosphorylated in a manner similar to that in oocyte chromosomes. In addition, our results show that the histone deacetylation and reacetylation of somatic chromosomes parallel the changes in oocyte chromosomes. Although the histone H3 phosphorylation and acetylation in somatic nuclei are reversible in oocyte cytoplasm, this methylation of H3–K9 in somatic nuclei is stable even after activation.


   Acknowledgements
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Acknowledgements
References
 
This work is supported, in part, by a Grant-in-Aid for Creative Scientific Research (13GS0008) to T.M. and by the 21st Century COE Program to T.M and H-T. B. from the Ministry of Education, Culture, Sports, Science and Technology of Japan. We thank the staff of the Kobe Meat Inspection Office facility for supplying the pig ovaries.


   Footnotes
 
First decision 23 August 2005
Received 21 July 2005
Accepted 7 February 2006
Revised manuscript received 31 January 2006


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Acknowledgements
 References
 

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