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Biology of Reproduction 65, 986-993 (2001)
© 2001 Society for the Study of Reproduction, Inc.

Regular Article

Posttranscriptional Regulation of Cyclin A1 and Cyclin A2 During Mouse Oocyte Meiotic Maturation and Preimplantation Development1

Dai-ichiro Fuchimotoa, Aki Mizukoshib, Richard M. Schultzc, Senkiti Sakaia, and Fugaku Aoki2,a,b

a Department of Animal Breeding, Graduate School of Life and Agricultural Science and b Department of Integrated Biosciences, Graduate School of Frontier Sciences, University of Tokyo, Bunkyo-ku, Tokyo 113-8657, Japan c Department of Biology, University of Pennsylvania, Philadelphia, Pennsylvania 19104-6018

ABSTRACT

A shift from a meiotic cell cycle to a mitotic cell cycle occurs following fertilization. The molecular basis for this transition, however, is poorly understood. Although cyclin A1 is proposed to regulate M phase in the meiotic cell cycle, and cyclin A2 is proposed to regulate S and M phases in the mitotic cell cycle, little is known about changes in the expression levels of cyclin A1 and A2 during meiotic and mitotic cell cycles in mammalian oocytes. We report that the mRNA levels of both cyclins A1 and A2 decrease during oocyte maturation. The amount of cyclin A1 mRNA then increases between the one-cell and blastocyst stages, whereas that of cyclin A2 remains relatively constant. The amount of cyclin A1 protein declines during maturation and is not readily detected from the two-cell to the blastocyst stage. In contrast, cyclin A2 is not readily detected in the oocyte and metaphase II-arrested egg but is detected following fertilization and throughout the subsequent stages of preimplantation development. The appearance of cyclin A2 protein following fertilization positively correlates with an increase in the size of the mRNA. This increase, as well as the increase in the amount of cyclin A2 protein, is prevented by 3'-deoxyadenosine (3'-dA), an inhibitor of polyadenylation. Consistent with a role for cyclin A2 in regulating the G1/S transition, 3'-dA also inhibits DNA replication in treated one-cell embryos. These results suggest that regulation of expression of cyclins A1 and A2 is under posttranscriptional regulation and that the observed changes in their expression may be involved in the transformation of a meiotic cell cycle to a mitotic cell cycle following fertilization.

developmental biology, early development, embryo, gamete biology, meiosis

INTRODUCTION

The appearance of an S phase characterizes one of the dramatic changes that occurs as oocytes of most species mature, are fertilized, and then undergo a series of reductive cleavage divisions. During mouse oocyte maturation, oocytes undergo a meiotic cell cycle in which progression from MI to MII occurs without an intervening S phase. Following fertilization, S phase occurs before M phase, which is typical of a mitotic cell cycle. This S phase differs, however, from that of a somatic cell in that DNA replication occurs separately in the two pronuclei, rather than in a common nucleus. In addition, G1 of the one-cell embryo is quite prolonged and the pronuclei do not form until some 5–6 h postfertilization. Once the pronuclei form, DNA replication rapidly ensues. Thus, in contrast to somatic cells, G1 in the one-cell embryo is characterized by a lack of transcription, which first initiates about midway through S phase [1]. The molecular basis for this transition in S phase regulation that occurs following fertilization is poorly understood.

Cyclin-dependent kinases (cdks) and cyclins are the key regulators of cell cycle progression [2–4]. Cdk2 and cdk4 are the primary regulators of commitment to enter S phase, and cdk1 (cdc2) is the primary regulator of entry into M phase. The activity of each cdk is regulated by its association with a specific cyclin. S-phase cyclins regulate the G1/S transition, and M-phase cyclins regulate the G2/M transition. The G1/S cyclins include cyclin D and cyclin E, which associate with cdk2 or cdk4. The G2/M cyclins include cyclin B, which associates with cdk1. In contrast to these cyclins that function during a single cell cycle transition, cyclin A regulates two phases; namely, progression through S phase and entry into M phase [5].

There are two subtypes of cyclin A, cyclin A1 and cyclin A2 [6–8]. In mammalian somatic cells cyclin A2 functions during both S and M phases [5, 9]. In contrast, meiotic cells (oocytes and spermatocytes) express cyclin A1 [7, 8]. This cyclin may function as the M-phase cyclin because the progression of spermatogenesis is arrested before the second meiosis in cyclin A1-deficient mice [10]. Thus, cyclin A1 is proposed to regulate M phase in the meiotic cell cycle, whereas cyclin A2 is proposed to regulate S and M phases in the mitotic cell cycle. These differences in the expression patterns of cyclin A1 and A2 may, therefore, be involved in the meiotic-mitotic transformation of cell cycle regulation after fertilization. However, there is no report that focuses on changes in the expression levels of cyclins A1 and A2 during meiotic and mitotic cell cycles in mammals.

In somatic cells, the expression of cyclin A2 is stringently regulated during progression of the mitotic cell cycle. Its expression abruptly increases just before S phase and decreases at M phase [11–15], and this change in expression is controlled essentially at the transcriptional level [16, 17]. Transcription does not occur, however, during the period following chromosome condensation and germinal vesicle breakdown (GVBD) in the maturing oocyte and in the early one-cell embryo prior to pronucleus formation [1, 18]. Maternal mRNAs that are recruited during oocyte maturation and following fertilization direct the changes that occur in gene expression during this developmental transition [19]. Thus, these fundamental differences in the regulation of gene expression at the transcriptional and posttranscriptional levels suggest that differences in the regulation of A-type cyclin expression should exist between oocytes/one-cell embryos and somatic cells. There is, however, little knowledge about the regulation of A-type cyclin expression in mouse oocytes and early embryos.

We report here that the expression of cyclin A1 and A2 protein in mouse oocytes and early embryos is not linked to their abundant mRNAs. Although the amount of cyclin A1 protein decreases between the oocyte and one-cell stages, the amount of cyclin A2 increases. The increase in the amount of cyclin A2 protein is coupled with polyadenylation of its mRNA. Treatment of one-cell embryos with 3'-deoxyadenosine (3'-dA) inhibits polyadenylation of cyclin A2 mRNA, the expression of cyclin A2 protein, and DNA synthesis. Thus, cyclin A2 mRNA appears to be a maternal mRNA that is recruited following fertilization and is responsible, at least in part, for S phase regulation in the one-cell embryo.

MATERIALS AND METHODS

Collection and Culture of Oocytes and Embryos

Female ddY mice 21–23 days of age and mature male ICR mice were purchased from SLC Japan (Shizuoka, Japan). Female mice were superovulated by injection with 5 IU of eCG. Full-grown oocytes were collected in Whitten medium [20] containing 0.2 mM 3-isobutyl-1-methylxanthine (IBMX) 45 h after eCG injection, and the cumulus cells were removed. The oocytes were then washed in IBMX-free medium and cultured at 38°C in an atmosphere of 5% CO2 and 95% air. The oocytes were observed 2 h after being transferred to IBMX-free medium and those that had not undergone GVBD were removed.

To obtain ovulated metaphase II-arrested eggs, eCG-primed mice were injected with 5 IU of hCG 48 h later. Ovulated eggs were collected from the oviducts 15–16 h after hCG injection. Sperm were collected from the cauda epididymis. The oocytes were inseminated with capacitated sperm, which had been incubated for 2 h at 38°C in an atmosphere of 5% CO2 and 95% air. Two hours after insemination, the embryos were washed with glucose-free CZB medium [21] and cultured at 38°C in an atmosphere of 5% CO2 and 95% air. The embryos were synchronized by removing those that had not yet formed a pronucleus 5.5 h after insemination. The embryos containing a pronucleus were transferred into CZB medium containing 5 mM glucose 48 h after insemination.

Semiquantitative Reverse Transcription-Polymerase Chain Reaction

Reverse transcription-polymerase chain reaction (RT-PCR) was used to quantify changes in the relative amount of mRNA [22]. Total RNA was isolated from either 100 oocytes or embryos by the acid guanidinium phenol-chloroform method and reverse-transcribed using SuperScript II and random hexamers (Gibco BRL, Grand Island, NY) according to the manufacturer's instructions. PCR was performed using a set of following primers:

  1. Cyclin A1:
  2. 5'-CTGTAGTTCTTCCCCTGCA-3' (sense)
  3. 5'-GCAAACAGCAAGTTGTTTATT-3' (antisense)
  4. Cyclin A2:
  5. 5'-GAGGTGGGAGAAGAATATAA-3' (sense)
  6. 5'-ACTAGGTGCTCCATTCTCAG-3' (antisense)
  7. Rabbit globin:
  8. 5'-GCAGCCACGGTGGCGAGTAT-3' (sense)
  9. 5'-GTGGGACAGGAGCTTGAAAT-3' (antisense)

The reaction was performed with either 28 cycles for cyclin A2 and rabbit globin or 35 cycles for cyclin A1. Each cycle consisted of 20 sec of denaturation at 94°C, 30 sec of annealing at either 55°C for cyclin A2 and rabbit globin or 58°C for cyclin A1 and 60 sec of extension at 72°C, followed by a final extension cycle at 72°C. The PCR products were separated by electrophoresis in 1.5% agarose, stained with ethidium bromide, and then photographed on Polaroid 665 negative/positive film. The negatives were scanned with a GS-670 densitometer and the value of the density of the band was determined with Molecular Analysis software (Bio-Rad, Hercules, CA). Preliminary experiments using known amounts of DNA indicated that a linear relationship was observed between the density of the band and the amount of DNA in a range of 3–90 ng.

For semiquantitative analysis, rabbit globin mRNA was added before isolation of total RNA. This served as an internal standard to evaluate the efficiency of RNA extraction and RT. PCR was conducted in the exponential range of amplification for each set of primers. The ranges in which the exponential amplification were observed were 30–36, 25–29, and 26–30 cycles for cyclin A1, cyclin A2, and globin, respectively. Therefore, PCR was performed by 35, 28, and 28 cycles for cyclin A1, cyclin A2, and globin, respectively. The ratio of the value of density for PCR products of cyclin A to globin was calculated.

Northern Blotting

Total RNA isolated from ~700 oocytes/embryos was separated on a 1.2% agarose gel containing 2.2 M formaldehyde, and transferred to a Hybond-N+ membrane (Amersham Life Science, Buckinghamshire, UK). The membrane was initially hybridized with salmon sperm DNA (Gibco BRL) at 65°C for 3 h and then hybridized with a digoxigenin (DIG)-labeled cRNA probe for cyclin A2 mRNA in ULTRAhyb buffer (Ambion, Austin, TX) at 65°C for 16 h. The DIG-labeled probe was synthesized by using a DIG RNA labeling kit (Boehringer-Mannheim, Mannheim, Germany) and pCR II plasmid vector (Invitrogen, Groningen, The Netherlands) inserted with 1269 base pairs (bp) of cyclin A2 sequence as a template. The hybridized probe was detected on x-ray film using the DIG Luminescent Detection kit (Boehringer-Mannheim).

Immunoblotting

Oocytes or embryos were lysed in 20 µl of extraction buffer (pH 7.4) composed of 50 mM Tris-HCl, 1% Nonidet P-40, 5 mM EGTA, 15 mM MgCl2, 2 mg/ml aprotinin, 2 mg/ml leupeptin, 1 mg/ml pepstatin, and 20 mg/ml phenylmethylsulfonyl fluoride. One hundred fifty oocytes and 70 embryos were used to detect cyclins A1 and A2, respectively. The samples were added to an equal volume of 2x SDS sample buffer [23] and boiled for 3 min. Proteins were separated on a 10% polyacrylamide gel and separated proteins were electrotransferred onto Immobilon-NC (Millipore, Bedford, MA). The membrane was blocked with Blocking Ace (Dainippon Pharmaceutical Co., Ltd., Osaka, Japan) for 30 min, and then incubated with the antibodies against cyclins A1 or A2, respectively (both C-terminal sequences from Santa Cruz Biotechnology, Inc., Santa Cruz, CA), in Tris-buffered saline containing 0.01% Tween-20 (TBST). In some experiments, antibody against whole cyclin A2 protein (Santa Cruz Biotechnology) was used. Unless otherwise specified, antibody against C-terminal peptide was used. The membrane was washed three times with TBST for 30 min each, followed by overnight incubation at 4°C with anti-goat immunoglobulin G (IgG; for cyclin A1) or anti-rabbit IgG (for cyclin A2) conjugated with horseradish peroxidase (Amersham Life Science) at a 1:2000 dilution. Immunoreactive proteins were then detected on x-ray film using a chemiluminescence reagent according to the manufacturer's instructions (NEN Life Science Products, Inc., Boston, MA). The amounts of proteins recognized by the antibodies were quantified by using a densitometer as described above.

Measurement of the Rate of Total Protein Synthesis

The embryos were metabolically rabiolabeled in medium containing 0.25 mCi/ml of [35S]methionine starting 6 h after insemination. Two hours later, embryos were washed six times with PBS containing 3 mg/ml polyvinylpyrrolidone and lysed in 40 µl of extraction buffer. The proteins were precipitated by adding 10 µl of 50 mg/ml BSA as a carrier and then 50 µl of 10% trichloroacetic acid. After centrifugation, the supernatant and pellet fractions were separated and the amount of radioactivity was determined by liquid scintillation counting. The results were expressed as the fraction of the number of counts per minute (cpm) in the pellet divided by the number of cpm in the pellet and supernatant.

Detection of DNA Synthesis

DNA synthesis was detected by the incorporation of bromodeoxyuridine (BrdU). Embryos were labeled with 10 µM BrdU from 7 to 8 h after insemination. The embryos were then washed in PBS containing 0.3% BSA (PBS/BSA), and next fixed with 3.7% paraformaldehyde. DNA was denatured by incubating the embryos in 2 N HCl at 37°C for 1 h. The acid was neutralized by incubating the samples in 0.1 M borate buffer pH 8.5 for 15 min. The incorporated BrdU was detected as previously described [24]. Briefly, the embryos were incubated with an anti-BrdU monoclonal antibody (Boehringer-Mannheim) for 45 min, followed by a 45-min incubation with an anti-mouse IgG antibody conjugated with Texas Red (Jackson ImmunoResearch Laboratories, West Grove, PA). The embryos were mounted on a glass slide with VectaShield (Vector Laboratories, Burlingame, CA) and observed with a laser-scanning confocal microscope.

RESULTS

Temporal Pattern of Cyclin A1 and A2 mRNA and Protein Expression

The changes in the amount of cyclin A1 and A2 mRNA were examined by semiquantitative RT-PCR during meiotic maturation and preimplantation development (Fig. 1). The relative abundance of both transcripts decreased during oocyte maturation. This is typical of many maternal mRNAs that are degraded following the onset of meiotic maturation. The relative abundance of the cyclin A1 transcript increased between the one-cell and two-cell stages, and continued to display a modest increase by the blastocyst stage. The increase in the relative abundance of the cyclin A1 mRNA was likely the result of zygotic transcription, which has definitely occurred by the two-cell stage. In contrast, cyclin A2 mRNA did not increase between the one-cell and two-cell stages, and moreover, the relative abundance remained essentially constant up to the morula stage; the relative abundance displayed a small increase at the blastocyst stage.



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  		FIG. 1. Temporal pattern cyclin A1 and A2 mRNA expression. The relative amounts of cyclin A1 and A2 transcripts were determined by RT-PCR in full-grown oocytes (GV), unfertilized eggs (E), one-cell embryos (1), two-cell embryos (2), four-cell embryos (4), morulae (M), and blastocysts (B). A) For cyclin A1 the value obtained in the blastocyst, which had the highest relative amount of transcript, was set as 100% and the values obtained for the other developmental stages were expressed relative to this value. B) For cyclin A2 the value obtained in the oocyte, which had the highest relative amount of transcript, was set as 100% and the values obtained for the other developmental stages were expressed relative to this value. The experiment was performed three times and the results are expressed as mean ± SEM

Peptide antibodies to cyclin A1 detected an immunoreactive band of Mr = 58 000 in germinal vesicle (GV)-intact oocytes (Fig. 2A). Staining was specific because the immunizing peptide, but not a peptide with an unrelated sequence, decreased the staining intensity. The amount of cyclin A1 in the metaphase II-arrested egg was less than that in the GV-intact oocyte, although its electrophoretic mobility was reduced (Fig. 3A). This retardation has been reported for Drosophila embryos in which two different migrating forms of cyclin A1 were detected [25]; the slower electrophoretic form could be due to phosphorylation. The decrease in cyclin A1 abundance was observed by 4 h following the initiation of oocyte maturation (i.e., subsequent to GVBD but prior to metaphase I (data not shown). Following fertilization, only the faster migrating form was observed (Fig. 3A). Although cyclin A1 was detected in the one-cell embryo 14 h postfertilization (cleavage occurs around 18 h postfertilization), little if any cyclin A1 was detected from the two-cell stage to the blastocyst stage. Thus, the amount of cyclin A1 in the oocytes and preimplantation embryos was uncoupled from the relative abundance of its transcript.



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  		FIG. 2. Specificity of antibodies used to detect cyclin A1 and cyclin A2. The antibody was initially incubated with antigen peptide (lane 2), control peptide with the sequence unrelated to cyclins A1 and A2 (lane 3), or no peptide (lane 1). A) GV-oocytes were used for the detection of cyclin A1. B) One-cell embryos were used for the detection of cyclin A2



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  		FIG. 3. Temporal pattern of cyclin A1 and A2 protein expression. Immunoblotting was conducted to determine the relative amounts of cyclin A1 (A) and cyclin A2 (B) at the same stages are described in the legend to Figure 1. The experiment was conducted two times and in which similar results were obtained

One-cell embryos also contained cyclin A2, as assessed by detection of two immunoreactive bands of Mr = 55 000 and 57 000 (Fig. 2B). The staining was specific because the immunizing peptide, but not a peptide with an unrelated sequence, decreased the staining intensity. The band with higher relative molecular mass probably corresponded to the phosphorylated form of cyclin A2, as is reported for Xenopus embryos [6]. It is interesting that cyclin A2 was not detected in GV-intact oocytes (Fig. 3B), which contained the highest abundance of cyclin A2 mRNA. Cyclin A2 staining was first detected in the one-cell embryo and was detected in all subsequent stages up to the blastocyst stage (Fig. 3B). Absence of cyclin A2 staining in GV-intact oocytes or metaphase II-arrested eggs and appearance in one-cell embryos were further confirmed by using the antibody against whole cyclin A2 protein (data not shown).

Cyclin A2 was readily detected 6 h following fertilization (Fig. 4), which is prior to the onset of S phase that occurs ~7 h postinsemination [24]. Recruitment of a maternal cyclin A2 mRNA following fertilization was the most likely explanation for the appearance of cyclin A2 protein within 6 h after insemination because there is no transcription in these embryos prior to S phase [1, 26]. Nevertheless, it was a formal possibility that a small amount of cyclin A2 mRNA was synthesized before S phase and that only this newly synthesized mRNA was efficiently translated, because the maternal mRNA apparently was not. Treatment of one-cell embryos with {alpha}-amanitin, an inhibitor of RNA polymerase II, did not prevent the appearance of cyclin A2 protein (data not shown). Thus, as predicted, the mobilization of a maternal cyclin A2 mRNA must account for the accumulation of cyclin A2 protein in G1 of the one-cell embryo.



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  		FIG. 4. Temporal pattern of cyclin A2 protein accumulation during oocyte maturation and following insemination. The oocytes and embryos were subjected to immunoblotting with anti-cyclin A2 antibody. The oocytes were collected at 0, 6, and 12 h after transfer to IBMX-free medium and the embryos were collected at 4, 6, 8, 11, and 14 h after insemination. Unfertilized eggs (E) were also collected and analyzed. The experiments were performed three times and a representative result is shown in A. The relative amounts of cyclin A2 protein were determined as described in Materials and Methods (B). The value obtained for embryos 14 h after insemination was set at 100% and values of other developmental stage were expressed relative to this value. The experiment was performed three times and the data are expressed as the mean ± SEM

Regulation of Cyclin A2 Protein Synthesis by Polyadenylation of Cyclin A2 mRNA

Elongation of the poly(A) tail of mRNAs is linked to the translational recruitment of many mRNAs in oocytes and embryos [19, 27, 28]. Accordingly, this posttranscriptional mode of regulation was examined for cyclin A2. Initial experiments using an RT-PCR approach [22, 29] to assess the elongation of the poly(A) tail were not successful for reasons that were not understood. Consequently, changes in mRNA length were analyzed by Northern blot analysis.

Northern blot analysis showed that two species of cyclin A2 mRNA differing in their length were expressed in the unfertilized eggs (Fig. 5); the two forms likely correspond to two alternatively spliced forms [30]. The length of both forms of cyclin A2 mRNA increased ~100 nucleotides by 6 h following insemination. Because no detectable level of mRNA synthesis occurs during this time, the longer form of each mRNA species could not be the product of de novo synthesis.



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  		FIG. 5. Northern blot analysis of cyclin A2 mRNA in unfertilized and fertilized eggs, and fertilized eggs treated with 3'-dA. A) Unfertilized oocytes were inseminated and then cultured with 2 mM 3'-dA from 1 h after insemination. The embryos cultured without 3'-dA were collected 0 (lane 1) and 8 h (lane 2), and those cultured with 3'-dA were collected 8 h after insemination. All samples were then processed for and subjected to Northern blot analysis. B) Densitometric tracing demonstrating the increase in the size of the cyclin A2 transcript following insemination and the inhibitory effect of 3'-dA on this increase. The experiment was performed three times and similar results were obtained in each case. Shown is a representative example. The upper, middle, and lower panels represent the results tracing lanes 1, 2, and 3, respectively, in A

Polyadenylation was the most likely explanation for the increase in size of the cyclin A2 transcripts. To confirm this, the embryos were treated with 3'-dA, which is metabolized to 3'-dATP and incorporated into the poly(A) tail during polyadenylation. Once 3'-dA is added to the poly(A) terminal, further addition of adenosine is inhibited [31–33]. Thus, 3'-dA functions as a chain terminator of poly(A) tail elongation. Northern blot analysis of one-cell embryos 8 h postinsemination revealed that the increase in the length of cyclin A2 mRNA was prevented by 3'-dA and that the size of the transcripts was essentially that observed in the control, uninseminated eggs (Fig. 5).

To examine whether the poly(A) tail elongation was involved in the accumulation of cyclin A2 protein after insemination, the embryos were treated with 3'-dA and examined for cyclin A2 protein. As anticipated, 3'-dA inhibited the accumulation of cyclin A2 protein by ~45%, when compared with the control treated with no reagent (Fig. 6). This inhibition was specific, because 3'-deoxyguanosine (3'-dG), a control for 3'-deoxypurine, or dimethylsulfoxide (DMSO), a control for solvent, had no inhibitory effect. In addition, the inhibitory effect of 3'-dA on cyclin A2 synthesis could not be attributed to a general inhibition of protein synthesis because the percentage of incorporation of [35S]methionine in the 3'-dA and control groups did not differ (58.2% ± 3.8% and 58.6% ± 3.4%, respectively).



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  		FIG. 6. Effect of 3'-dA on cyclin A2 accumulation. Embryos were treated with 2 mM 3'-dA, 2 mM 3'-deoxyguanosine (dG), 0.4% DMSO, or no reagent (cont.) from 1 h after insemination. The 3'-deoxypurines were dissolved in DMSO to make the stock solution and the final concentration of DMSO was 0.4% when 3-deoxypurines were added to the culture medium. The embryos were collected for immunoblotting with anti-cyclin A2 antibody 8 h after insemination. The experiment was conducted five times (except for 3'-dG, which was conducted four times) and a representative result is shown in A. The relative amounts of proteins were quantified as described in Materials and Methods (B). The value obtained for embryos cultured with no reagent (cont.) was set as 100% and values for the embryos cultured with the reagents were expressed relative to this value. The data are expressed as the mean ± SEM. The value of 3'-dA was significantly different from those of 3'-dG and DMSO (P < 0.05, by the Student t-test)

Inhibitory Effect of 3'-dA on DNA Synthesis

Cyclin A2 is required for DNA synthesis in somatic cells [13, 34]. Results from the studies described above suggest that the appearance of cyclin A2 prior to S phase in the one-cell embryo could be essential for initiating S phase. Accordingly, inhibiting cyclin A2 accumulation by treating one-cell embryos with 3'-dA would be expected to inhibit the onset of DNA replication in these treated embryos. Embryos were cultured with 3'-dA and BrdU incorporation was examined 8 h postinsemination. In the control embryos, at 8 h after insemination >80% of female pronuclei and almost all male pronuclei had incorporated BrdU. In contrast, treatment with 3'-dA decreased the percentages of pronuclei incorporating BrdU in a concentration-dependent manner (Fig. 7). This inhibition was specific because neither 3'-dG nor DMSO resulted in any significant inhibition. Thus, as anticipated, 3'-dA inhibited DNA replication.



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  		FIG. 7. Effect of 3'-deoxyadenosine (3'-dA) on DNA synthesis in one-cell embryos. The embryos were cultured with 0.5, 2, or 5 mM 3'-dA, 2 mM 3'-deoxyguanosine (dG), 0.4% DMSO (G), or no reagent (C) from 1 h after insemination. The 3'-deoxypurines were dissolved in DMSO to make the stock solution and the final concentration of DMSO was 0.4% when 3-deoxypurines were added to the culture medium. Five millimoles of 3'-dG could not be prepared because it did not dissolve completely in DMSO. The embryos were examined for DNA synthesis as described in Materials and Methods 8 h after insemination. Open and closed columns show the percentages of the male and female pronuclei incorporating BrdU, respectively. The experiment was performed four times, similar results were obtained in each case, and data were pooled. The numbers of embryos examined were 59 (C), 117 (D), 68 (0.5 mM 3'-dA), 119 (2 mM 3'-dA), 60 (5 mM 3'-dA), and 136 (2 mM 3'-dG)

DISCUSSION

We report here that the temporal patterns of expression of cyclin A2 at the mRNA and protein levels are unlinked in mouse oocytes and preimplantation embryos (Figs. 1 and 3). Moreover, whereas cyclin A1 is present in oocytes and one-cell embryos it is not detected from the two-cell stage onward (Fig. 3A). In contrast, cyclin A2 is not readily detected in the oocyte but is readily detected in the one-cell embryo and remains throughout preimplantation development (Fig. 3B). Thus, cyclin A1 is preferentially expressed during the meiotic cell cycle, whereas cyclin A2 is preferentially expressed during the mitotic cell cycle. Mobilization of a maternal cyclin A2 mRNA following fertilization appears to be responsible for the dramatic increase in the amount of cyclin A2 in the one-cell embryo in comparison to the oocyte (Figs. 4–6), and inhibiting this mRNA recruitment inhibits entry into S phase of the one-cell embryo (Fig. 7). While our studies were in progress, a report appeared that also noted that in the mouse the amount of cyclin A2 protein increases between the oocyte and one-cell stage [35]; the differences in the magnitude of the changes likely reflect differences in the sensitivity of the antibodies used.

The expression patterns of A-type cyclins in Xenopus differ from that in the mouse. In Xenopus, cyclin A1 is expressed during the meiotic cell cycle as in mouse, but persists up to the blastula stage [6, 36], at which time expression of cyclin A2 commences [6]. This difference may reflect differences in the developmental stage at which maternal-zygotic transition of developmental control occurs. Howe et al. [6] suggested that the mechanism controlling the abrupt decrease in cyclin A1 and increase in A2 at the blastula stage is dependent on time of development, but not on cell division or transcription, and that this mechanism plays an essential role in releasing embryos from maternal control in Xenopus. This hypothesis is consistent with the present results. In mouse, the maternal-to-embryonic transition occurs at the two-cell stage [18]. Alternatively, the difference of the expression pattern of A-type cyclins between Xenopus and mouse may reflect the difference in the acquisition of checkpoint control mechanisms during development. In Xenopus, prior to the blastula stage, which is reached within 7 h following fertilization, the embryos divide rapidly and the 12 cell cycles consist of successive S and M phases, without an intervening G1 or G2 phase. In contrast, in the mouse, cell cycle times are not rapid (e.g., the first cell cycle takes about 20 h [37]), and a bona fide cycle exists (i.e., G1, S, G2, and M phases). Thus, Xenopus embryos apparently lack functional checkpoints prior to the blastula stage. In fact, acquisition of functional checkpoints monitoring the completion of S phase and entry into M phase are different between these two species. One-cell mouse embryos do not enter M phase when DNA synthesis is inhibited [38], indicating that the checkpoint is functional shortly after fertilization. In contrast, Xenopus embryos enter M phase up to the blastula stage even if DNA synthesis is inhibited [39], indicating that this checkpoint does not operate until the blastula stage. Thus, in each of these species, the onset of a functional G2/M checkpoint coincides with the expression of cyclin A2 protein. In Xenopus, the lack of functional checkpoints to monitor cell cycle progression may be a consequence of the rapid cell divisions prior to the mid-blastula transition, because a functional checking mechanism could constrain the rapid development that is intrinsic to this species. In contrast, in the mouse, cell cycle times are not rapid and the bona fide cycle likely requires the presence of a functional checking mechanism.

Only one type of cyclin A is found in marine invertebrates and Drosophila, and it is expressed in both meiotic and mitotic cell cycles [25, 40, 41]. Phylogenetic tree analysis using nucleotide sequence suggests that the A-type cyclin in these animals is a prototype of cyclin A1 and A2 in vertebrates and that this prototype is generated from cyclin B (Fig. 8). This evolution may reflect a role for cyclin A2 in the checking mechanism for cell cycle progression, as described above, that evolved as the complexity or organisms increased. In fact, inhibiting cyclin A-dependent kinase in S-phase prevents cells from entering M phase [42]. The appearance of significantly higher amounts of cyclin A2 following fertilization would permit a functional M-phase checkpoint to develop following fertilization, whereas the presence of cyclin A1 may regulate the meiotic cycle up to arrest at metaphase II.



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  		FIG. 8. Phylogenetic tree of cyclins. Amino acid sequences of A) cyclin family members in mouse and B) A-type cyclins in various species were analyzed by CLASTALW (National Institute of Genetics, available at http://www.ddbj.nig.ac.jp/E-mail/clustalw-j.html) and expressed by TreeView (http://taxonomy.zoology.gla.ac.uk/rod/rod.html)

Elongation of the poly(A) tail has been proposed to regulate the translational control of maternal mRNAs during meiotic maturation and early preimplantation development [20], although some other mechanisms may also be involved (e.g., binding of masking proteins to the 3' untranslated region; UTR) [43, 44]. There is controversy, however, about the role of poly(A) elongation. Although some reports suggest that poly(A) tail elongation itself does not directly cause the increase in the efficiency of translation [45], other reports show that the change in poly(A) length is tightly correlated to the translational efficiency of the mRNA. In mouse oocytes, poly(A) tail elongation is reported to regulate the synthesis of several proteins (i.e., tissue plasminogen activator [46], hypoxanthine phosphoribosyltransferase [47], and Mos protein [48]). There is, however, a paucity of information after regarding the role of poly(A) tail elongation in the recruitment of maternal mRNAs following fertilization of mouse eggs. Fertilization of mouse eggs is accompanied by changes in the pattern of protein synthesis. These changes occur within 4 h and hence are the manifestation of the recruitment of maternal mRNAs that are mobilized [49]. Only a few maternal mRNAs that are mobilized following fertilization have been identified; namely, an unidentified mRNA [50], and ß-catenin, Ptp4a1, and Spin [51]. Results presented here suggest that cyclin A2 is another such mRNA. The aforementioned mRNAs contain a polyuridine tract in the 3' UTR that is within ~1000 nucleotides from the polyadenylation signal sequence (AAUAAA). This polyuridine tract is proposed to serve as a cytoplasmic polyadenylation element (CPE) in Xenopus embryos [45, 52], whereas the (A)UUU(UU)AU(AA) consensus sequence [51, 53, 54] defines a CPE element that is used to recruit mRNAs during oocyte maturation (e.g., tPA [AUUUUAAU] and ß-actin [UUUUUAAU]). Examination of the cDNA sequence of the mouse cyclin A2 does not reveal an apparent polyuridine tract. Thus, there may be a more complicated "code" of 3' UTR sequences that direct the temporal recruitment of mRNAs during maturation and following fertilization.

Inhibiting poly(A) tail elongation by 3'-dA inhibits the synthesis of cyclin A2 and DNA replication in one-cell embryos, suggesting that cyclin A2 is involved in initiating DNA replication. Because 3'-dA would inhibit the mobilization of all maternal mRNAs, it is formally possible that the mobilization of mRNAs other than cyclin A2 mRNA are involved in initiating DNA replication in the one-cell embryo. Nevertheless, the observed inhibition of DNA replication is consistent with the requirement in somatic cells for cyclin A2 in DNA replication [12, 34]. It should be noted that Winston et al. [35] showed that cyclin A2-null mutant embryos develop normally from the four-cell stage to the postimplantation stage in the absence of detectable cyclin A2 protein. In mutant embryos, maternal cyclin A2 mRNA is degraded at the two- to four-cell transition and the protein is degraded at the second mitosis. Thus, cyclin A2 protein is present only in one- and two-cell embryos in the mutant mouse. Although these results clearly demonstrate that cyclin A2 is dispensable for preimplantation development after the two-cell stage, they shed no light on its role in one-cell and two-cell embryos. The results presented here suggest that cyclin A2 transcribed from maternal mRNA plays a role in DNA replication in the one-cell embryo. This role, however, may be lost following the maternal-to-embryonic transition and assumed by another, as yet to be identified, cyclin.

In summary, results presented here suggest that polyadendylation of cyclin A2 mRNA is linked to the recruitment of this maternal mRNA following fertilization. The marked increase in the amount of cyclin A2 protein that results from this mobilization may function in the conversion of a meiotic cell cycle, without an S phase, prior to fertilization to a mitotic one, with an S phase, postfertilization.

FOOTNOTES

First decision: 11 April 2001.

1 This research was supported by a grant from the Ministry of Education, Science and Culture to F.A. and grant HD22681 from the National Institutes of Health to R.M.S. Back

2 Correspondence: Fugaku Aoki, Department of Animal Breeding, University of Tokyo, Yayoi 1-1-1, Bunkyo-ku, Tokyo 113-8657, Japan. FAX: 81 3 5841 8180; aokif{at}k.u-tokyo.ac.jp Back

Accepted: May 7, 2001.

Received: March 7, 2001.

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