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Dependence of DNA Synthesis and In Vitro Development of Bovine Nuclear Transfer Embryos on the Stage of the Cell Cycle of Donor Cells and Recipient Cytoplasts¹

^a Laboratory of Reproductive Biology, Graduate School of Agriculture, Kyoto University, Kyoto 606-8502, Japan

	ABSTRACT

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The effect of the stage of the cell cycle of donor cells and recipient cytoplasts on the timing of DNA replication and the developmental ability in vitro of bovine nuclear transfer embryos was examined. Embryos were reconstructed by fusing somatic cells with unactivated recipient cytoplasts or with recipient cytoplasts that were activated 2 h before fusion. Regardless of whether recipient cytoplasts were unactivated or activated, the embryos that were reconstructed from donor cells at the G0 phase initiated DNA synthesis at 6–9 h postfusion (hpf). The timing of DNA synthesis was similar to that of parthenogenetic embryos, and was earlier than that of the G0 cells in cell culture condition. Most embryos that were reconstructed from donor cells at the G1/S phase initiated DNA synthesis within 6 hpf. The developmental rate of embryos reconstructed by a combination of G1/S cells and activated cytoplasts was higher than the rates of embryos in the other combination of donor cells and recipient cytoplasts. The results suggest that the initial DNA synthesis of nuclear transfer embryos is affected by the state of the recipient oocytes, and that the timing of initiation of the DNA synthesis depends on the donor cell cycle. Our results also suggest that the cell cycles of somatic cells synchronized in the G1/S phase and activated cytoplasts of recipient oocytes are well coordinated after nuclear transfer, resulting in high developmental rates of nuclear transfer embryos to the blastocyst stage in vitro.

developmental biology, early development, embryo

	INTRODUCTION

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Viable cloned animals from somatic cells have been produced by nuclear transfer techniques in sheep [1], cattle [2–4], mice [5], goats [6], and pigs [7, 8]. However, the efficiency of cloning has been extremely low, and factors involving the efficiency are still unclear.

Unactivated oocytes have been generally used as recipient cytoplasts for nuclear transfer of somatic cells, because of the poor developmental ability of nuclear transfer embryos reconstructed from activated recipient cytoplasts [9–11]. However, it was reported that cytoplasts that were prepared by enucleation of chromosomes at telophase after activation of oocytes in cattle [12–14] or by chemical enucleation with demecolcine after activation in mice [15] are useful for producing cloned animals from somatic cells. Therefore, not only unactivated recipient cytoplasts but also activated cytoplasts may support a reprogramming of somatic cell nuclei.

In the first report of production of a live offspring from somatic cells, donor cells were synchronized at the G0 phase by serum starvation [1]. This result suggests that the chromatin at the G0 phase is amenable to nuclear reprogramming [16]. However, donor cells in phases other than the G0 phase can be used to produce cloned animals [2, 11], indicating that synchronization in the G0 phase is not a prerequisite for somatic cloning. In cattle, when unactivated cytoplasts were used as the recipient, embryos that were reconstructed from donor cells at phases of the cell cycle other than the S phase were able to develop to the blastocyst stage, but embryos that were reconstructed from donor cells at the S phase did not develop to the blastocyst stage [11]. In mice, nuclear transfer embryos that were reconstructed from unactivated recipient cytoplasts and donor cells at the M phase showed a higher developmental ability than those reconstructed from unactivated recipient cytoplasts and donor cells at interphase [17]. These studies indicate that the developmental ability of nuclear transfer embryos depends on both the cell cycle stage of the donor cells and the cytoplasmic state of the recipient oocytes.

The development of nuclear transfer embryos, besides depending on the synchronization of the cell cycles of the donor cells and recipient cytoplasts at the time of nuclear transfer, may also depend on aspects of the embryos after nuclear transfer, such as the timing of entrance into the S phase, the duration of the S phase, and the timing of first mitosis. However, the effect of combination of the cell cycle of donor cells and the cytoplasmic state of recipient oocytes on the progression of cell cycle has not been reported.

In this study, we examined how the cell states of the donor cells and the recipient cytoplasts affect the in vitro development of nuclear transfer embryos, and the cell cycle progression of the embryos after nuclear transfer.

	MATERIALS AND METHODS

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Preparation of Donor Cells

Bovine ear skin cells from an adult Holstein cow were cultured in Dulbecco modified Eagle medium (DMEM; Invitrogen, Carlsbad, CA) supplemented with 10% fetal bovine serum (FBS; JRH Biosciences, Lenexa, KS), and used as untreated donor cells for nuclear transfer at 80% to 90% of confluency. For cell-cycle synchronization of donor cells, the cells were further cultured in DMEM supplemented with 0.2% FBS for 2 days. After serum starvation for 2 days, cells were passaged and separated into two groups. One group of cells was synchronized at the G0 phase by culturing them in DMEM supplemented with 0.2% FBS for 20 h. The other group of cells was synchronized at the G1/S phase by culturing them in DMEM supplemented with 10% FBS and 2 µg/ml of aphidicolin (Sigma, St. Louis, MO) for 20 h.

Preparation of Recipient Cytoplasts

Ovaries were obtained from a slaughterhouse and brought to the laboratory in saline at 39°C. Cumulus-oocyte complexes (COCs) were aspirated from follicles with diameters within 8 mm using a 21-gauge needle and washed in a maturation medium. The maturation medium was tissue culture medium-199 (TCM-199; Nissui, Tokyo, Japan) supplemented with 10% FBS, 100 IU/ml of hCG (Puberogen; Sankyo, Tokyo, Japan) and 1 µg/ml of estradiol-17ß. The COCs were placed in droplets of maturation medium and cultured for 18–20 h at 39°C under an atmosphere of 5% CO₂ in air. After in vitro maturation, the cumulus cells were removed by vortexing COCs in Hepes-buffered TCM-199 supplemented with 0.1% hyaluronidase (Sigma) for 4 min. To enucleate oocytes, a small volume of cytoplasm (about 25%) laying beneath the first polar body was squeezed out of the zona pellucida using a glass needle [10]. The removed cytoplasm was stained with Hoechst 33258 (Sigma) and examined under UV light to confirm that it contained the metaphase chromosomes. Enucleated cytoplasts were randomly assigned to two different nuclear transfer protocols described below.

Nuclear Transfer

Nuclear transfer was performed by two protocols (Fig. 1). In one protocol, recipient cytoplasts were fused with donor cells and simultaneously activated at 24 h postmaturation (hpm) by DC electric pulses at 200 V/mm for 2 x 10 µsec, and then treated with 10 µg/ml of cycloheximide (Sigma) for 6 h (F24). The second protocol was the same as the first except that 2 h before fusion, the recipient cytoplasts were activated with 7% ethanol for 7 min and then treated with cycloheximide for 2 h (A22F24). Nuclear transfer embryos were cultured in synthetic oviduct fluid (SOF) [18] supplemented with 2% Eagle basal medium (BME) amino acids solution (Sigma), 1% modified Eagle medium (MEM) nonessential amino acids solution (Sigma) and 1% FBS. At 2 days after fusion, the cleaved embryos were counted and then transferred to SOF supplemented with 2% BME amino acids solution, 1% MEM nonessential amino acids solution, and 5% FBS. The transferred embryos were cultured for an additional 5 days, and the developmental rates of embryos to the blastocyst stage were examined.

View larger version (12K): [in this window] [in a new window]   FIG. 1. Protocol and time course of nuclear transfer

Western Blotting Analysis

Recipient cytoplasts were collected in SDS sample buffer containing 62.5 mM Tris-HCl (pH 6.8), 10% glycerol (Wako Pure Chemical Industries, Osaka, Japan), 20 mg/ml of SDS (lauryl sulfate; Sigma) and 1% 2-mercaptoethanol, and boiled for 3 min to detect cyclin B1 by a Western blotting analysis. The samples were separated by 10% PAGE and transferred to a cellulose nitrate membrane. The membrane was blocked by 5% skim milk in PBS containing 0.1% Tween 20 (PBS-T), washed three times in PBS-T, and incubated with cyclin B1 antibody (sc-595, Santa Cruz Biotechnology, Santa Cruz, CA) diluted 1:250 with PBS-T for 4 h at room temperature. After three washes in PBS-T, the membrane was incubated with biotinylated swine anti-rabbit immunoglobulins (DAKO, Glostrup, Denmark) diluted by 1:6000 with PBS-T for 1 h at room temperature. After three washes in PBS-T, the membrane was incubated with horseradish peroxidase-conjugated streptavidin (Amersham Pharmacia Biotech, Buckinghamshire, U.K.) diluted 1:1500 with PBS-T for 1 h at room temperature. The membrane was washed three times in PBS-T and then the blot was visualized with enhanced chemiluminescence (Amersham).

Detection of DNA Synthesis of Synchronized Donor Cells and Nuclear Transfer Embryos

DNA synthesis was detected by an immunofluorescence assay using a 5-bromo-2'-deoxyuridine (BrdU) Labeling and Detection Kit I (1 296 736l; Roche Diagnostics, Indianapolis, IN). To examine the timing of DNA synthesis of donor cells, cells were synchronized in the G0 or G1/S phase as described above, and after being released from synchronization treatment, DNA synthesis was detected according to the manufacturer's procedure. To examine the timing of DNA synthesis of nuclear transfer embryos, embryos were labeled with BrdU in a medium during 3–6 h postfusion (hpf) or 6–9 hpf, and BrdU uptake was examined at 6 hpf or 9 hpf, respectively. Labeled embryos were fixed on coverslips, permeabilized, and hydrolyzed according to the method described by Campbell et al. [19]. After hydrolysis, the embryos were treated with anti-BrdU solution, and then with anti-mouse immunoglobulin-fluorescein solution according to the manufacturer's procedure. Samples were mounted on glass slides and observed under a fluorescent microscopy. To compare the timing of DNA synthesis between nuclear transfer embryos and parthenogenetic embryos, matured oocytes were parthenogenetically activated with 7% ethanol for 7 min followed by cycloheximide treatment for 6 h, and the timing of DNA synthesis was examined.

Statistical Analysis

Data were analyzed by the Tukey method using the general-linear model procedure of the Statistical Analysis System (SAS Institute, Cary, NC).

	RESULTS

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Inactivation of Maturation Promoting Factor in Activated Recipient Cytoplasts

Expression of cyclin B1, one of the components of maturation-promoting factor (MPF), in recipient cytoplasts used in the F24 and A22F24 protocols was examined by a Western blotting analysis (Fig. 2). Expression of cyclin B1 was detected in F24 cytoplasts, indicating that F24 cytoplasts were arrested at metaphase II (MII) with high MPF activity. In A22F24 cytoplasts, cyclin B1 was not detected, and MPF had already lost its activity.

View larger version (21K): [in this window] [in a new window]   FIG. 2. Expression of cyclin B1 in an unactivated cytoplast (F24) and in a preactivated cytoplast (A22F24) 2 h after activation

Development of Nuclear Transfer Embryos Using Untreated Donor Cells

Activated cytoplasts with low or no MPF activity can support in vitro development of nuclear transfer embryos as well as unactivated cytoplasts with high MPF activity (Table 1). The rates of development to the blastocyst stage were not significantly different between F24 and A22F24.

Timing of DNA Synthesis of Synchronized Donor Cells

The timing of DNA synthesis of donor cells after being released from serum starvation or aphidicolin treatment was examined (Table 2). In cells synchronized in the G0 phase by serum starvation, DNA synthesis did not occur until 12 h after the end of starvation. In cells synchronized at the G1/S phase by aphidicolin, 66.7% of cells started DNA synthesis at 1 h after aphidicolin treatment and more than 80% of cells entered S phase by 3 h.

Timing of DNA Synthesis of Nuclear Transfer Embryos

Nuclear transfer embryos that were reconstructed from G1/S donor cells initiated DNA synthesis earlier than those reconstructed from G0 cells (Table 3). None of the parthenogenetic embryos initiated DNA synthesis within 6 h after activation. Also, only a few of the nuclear transfer embryos that were reconstructed from G0 donor cells initiated DNA synthesis within 6 hpf. On the other hand, more than 60% of embryos that were reconstructed from G1/S cells initiated DNA synthesis within 6 hpf. At 9 hpf, most of the embryos had initiated DNA synthesis regardless of the stage of the cell cycle of the donor cells or the timing of nuclear transfer.

Development of Nuclear Transfer Embryos Using Synchronized Donor Cells

In vitro development of embryos that were reconstructed from synchronized donor cells (G0 or G1/S) by fusion with unactivated (F24) or preactivated (A22F24) cytoplasts was examined (Table 4). The cleavage rate of embryos reconstructed from A22F24 cytoplasts was significantly higher than the rate of embryos reconstructed from F24 cytoplasts. The highest cleavage rate and developmental rate were recorded in A22F24-G1/S embryos. The rate of development of A22F24-G1/S embryos to the blastocyst stages was significantly higher than the rates of F24-G1/S and A22F24-G0 embryos, and was higher than that of F24-G0 embryos without a significant difference.

	DISCUSSION

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Unactivated oocytes have been generally used as recipient cytoplasts for production of cloned animals from somatic cells [1, 2, 4, 5, 7, 8]. Rates of development to the blastocyst stage in vitro of somatic nuclear transfer embryos that were reconstructed from activated cytoplasts were very low, suggesting that exposure of donor nuclei to unactivated recipient cytoplasts is beneficial or essential for reprogramming of somatic nuclei [9–11]. In this study, activated cytoplasts supported the development of somatic nuclear transfer embryos in vitro at rates comparable to those of embryos that were reconstructed from unactivated cytoplasts. The expression of cyclin B1, one of the components of MPF, was not detected at 2 h after activation (Fig. 2), indicating that MPF was inactivated in the activated cytoplasts used in this study. Vignon et al. [3] reported that by using recipient cytoplasts in a low MPF state induced by aging and cooling of enucleated oocytes, nuclear transfer embryos that were reconstructed from donor cells from calf or fetus developed to the blastocyst stage and to term [3]. Together, these data suggest that a high MPF activity in the recipient cytoplasts is not essential for the development of somatic nuclear transfer embryos, and that factors essential for a reprogramming of somatic nuclei exist in recipient cytoplasts at 2 h after activation. In cattle, recipient cytoplasts that were enucleated after activation, at the telophase stage, supported the development of somatic nuclear transfer embryos as well as unactivated recipient cytoplasts [13, 14]. In mice, recipient cytoplasts that were chemically enucleated by demecolcine after activation led to the development of nuclear transfer embryos to term at a high rate [15]. These studies and our results indicate that activated cytoplasts can be used for nuclear transfer as recipients. In contrast to the methods used in the previous studies, recipient cytoplasts used in our methods were enucleated before activation, suggesting that the developmental ability of somatic nuclear transfer embryos is not affected by the presence or absence of chromosomes in recipient cytoplasts at the time of activation.

The stage of the cell cycle of donor cells at the time of fusion with recipient cytoplasts affected the timing of DNA synthesis of nuclear transfer embryos (Table 3). The timing of initiation of DNA synthesis of embryos that were reconstructed from G0 cells was slower than that of embryos from G1/S cells regardless of the state of the recipient cytoplasts. Most of the donor cells that were synchronized at the G0 phase did not initiate DNA synthesis until 6 h after nuclear transfer. The timing of DNA synthesis was earlier than that of cells synchronized at G0 phase in culture and was similar to that of parthenogenetic embryos observed in this study. Previous studies [20, 21] have also reported that DNA synthesis of parthenogenetic embryos occurs after being released from protein synthesis inhibitors or cyclin-dependent kinases. Two possible explanations can be made from these results. First, one or more cytoplasmic components existing in recipient cytoplasts accelerate the cell cycle of G0 nuclei, enabling them to initiate DNA synthesis at 6–9 hpf; and second, the cell cycle of G0 nuclei progresses according to the cell cycle progression of recipient cytoplasts and DNA synthesis initiates after being released from a protein inhibitor (i.e., cycloheximide). On the other hand, most of the donor cells that were synchronized at the G1/S phase initiated DNA synthesis within 6 h after nuclear transfer regardless of whether recipient cytoplasts were unactivated or activated, suggesting that the donor nuclei at the G1/S phase are ready to enter the S phase after fusion within 6 hpf regardless of the presence of cycloheximide.

In this study, when donor cells were prepared without synchronization treatment, the cleavage rates of reconstructed embryos did not differ between F24 and A22F24 protocols. In contrast, when donor cells synchronized at the G0 or G1/S phase were used, the cleavage rate of embryos that were reconstructed by the A22F24 protocol was significantly higher than that of embryos reconstructed by the F24 protocol. A possible explanation of these results is that the chromatin structure of nuclei at phases of the cell cycle other than the M phase are damaged by transfer into unactivated cytoplasts at meiotic metaphase, and that the nuclei that are synchronized by serum starvation or chemical treatment are more sensitive to chromosomal damage. This hypothesis is compatible with a recent report that injection of nuclei at the M phase into unactivated cytoplasts provided a significantly higher developmental rate in vitro than injection of nuclei at interphase [17].

The stages of the cell cycle of the donor cells and the activation status of the recipient cytoplasts affected the ability of nuclear transfer embryos to develop to the blastocyst stage (Table 4). The developmental rate of A22F24-G1/S embryos was significantly higher than the rate of A22F24-G0 and F24-G1/S embryos. Although it was also higher than the developmental rate of F24-G0 embryos, the differences were not significant. As described above, embryos that were reconstructed from donor cells synchronized at the G1/S phase enter the S phase earlier than those reconstructed from donor cells synchronized at the G0 phase. Because chromatin unfolds during DNA synthesis to make it more accessible [22], we presume that the earlier entrance of transferred nuclei into S phase allows reprogramming factors to easily access the donor chromatin before they disappear. Taken together, these data suggest that donor cells at the G1/S phase, in combination with A22F24 cytoplasts, are better synchronized with each other at the time of fusion and during subsequent cell cycle progression than are other combinations.

In conclusion, our results indicate that the cell cycle progression of nuclear transfer embryos is affected by recipient cytoplasts in the manner that depends on the cell cycle of donor cells at the time of nuclear transfer, and that donor cells synchronized at the G1/S phase may be suitable for recipient cytoplasts activated 2 h before fusion in bovine somatic cell nuclear transfer.

	ACKNOWLEDGMENTS

 
We thank M. Goto and H. Kasai of the Livestock Research Institute in Tokushima Prefecture for providing somatic cells, and A. Ohashi of Kyoto University for his technical advice on Western blotting analysis.

	FOOTNOTES

 
First decision: 21 January 2002.

¹ This work was supported by The Japan Society for the Promotion of Science, "Research for the Future" Program (JSPS-RFTF97 L00905) to H.I.

² Correspondence. FAX: 81 75 753 6329; imai{at}kais.kyoto-u.ac.jp

Accepted: March 18, 2002.

Received: December 27, 2001.

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