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. 2002 Jan;14(1):197–210. doi: 10.1105/tpc.010386

The Rice Cyclin-Dependent Kinase –Activating Kinase R2 Regulates S-Phase Progression

Tanja Fabian-Marwedel a, Masaaki Umeda b, Margret Sauter a,1
PMCID: PMC150560  PMID: 11826308

Abstract

Cyclin-dependent kinases (CDKs) are the central components of eukaryotic cell cycle regulation. Phosphorylation of CDKs at a conserved threonine residue is required for their full activity and is mediated by a CDK-activating kinase (CAK). The CAK R2 from rice belongs to those CAKs that phosphorylate not only CDKs but also the C-terminal domain (CTD) of RNA polymerase II. We showed that R2 is a nuclear protein with increased expression and increased CTD kinase activity in S-phase. Increasing R2 abundance through a transgenic approach accelerated S-phase progression and overall growth rate in suspension cells. In planta, the CTD kinase activity of R2 was induced by a growth-promoting signal. R2 regulation, therefore, may constitute a plant-specific adaptive mechanism that is used to adjust the rate of cell proliferation in response to a changing environment.

INTRODUCTION

Cell division is regulated by the sequential activation of cyclin-dependent kinases (CDKs). CDKs are among the most highly regulated enzymes known. Their activities depend on binding to a regulatory cyclin subunit, binding to inhibitory subunits, subcellular localization, protein degradation, and multiple phosphorylation events (Morgan, 1995; Pines, 1995; Sherr and Roberts, 1995; King et al., 1996; Solomon and Kaldis, 1998). The kinase subunit by itself is inactive and requires binding to a cyclin and phosphorylation on a conserved threonine residue in the T-loop (e.g., Thr-161 in rice Cdc2Os-1) for full activation (Gould et al., 1991; Desai et al., 1992; Solomon et al., 1992; Yamaguchi et al., 1998). The kinase responsible for this phosphorylation has been termed CAK for CDK-activating kinase. CAKs are related to but clearly distinct from other classes of CDKs (Kaldis, 1999). Because phosphorylation of CDKs is a crucial step in their activation, much effort has been directed toward characterizing CAKs (Morgan, 1995).

In animals and yeast, two classes of CAKs have been identified. They are represented by human p40MO15/Cdk7 and by Cak1pCiv1 from budding yeast. These CAKs have low homol-ogy with each other, and they differ in their enzyme characteristics. Human Cdk7 has been shown to phosphorylate CDK substrates and the C-terminal domain (CTD) of the large subunit of RNA polymerase II, the second reaction as part of the general transcription factor IIH (Buck et al., 1995; Damagnez et al., 1995; Draetta, 1997). Because transcription factor IIH also is involved in nucleotide excision repair, Cdk7 may have functions in cell cycle control, in transcriptional regulation, and in DNA repair (Roy et al., 1994; Serizawa et al., 1995; Shiekhattar et al., 1995; Orphanides et al., 1996; DeLaat et al., 1999; Kaldis, 1999). Budding yeast Cak1p acts as a CAK but does not display CTD kinase activity.

Most CDKs are thought to be activated by CAKs, and CAKs are essential genes. Animal and yeast CAKs are expressed and active throughout the mitotic cell cycle with no change in subcellular localization, indicating that CAKs are not regulated in these organisms (Matsuoka et al., 1994; Poon et al., 1994; Tassan et al., 1994, 1995; Espinoza et al., 1996; Sutton and Freiman, 1997). CAK overexpression increased CAK activity but did not produce a mutant phenotype (Kaldis et al., 1996). Yet, CDK phosphorylation appears to have specific functions in cell cycle regulation. Immunodepletion of Xenopus XlCdk7 in egg extracts suppressed CAK activity and arrested cells before M-phase (Fesquet et al., 1997). Furthermore, Drosophila DmCdk7 was shown to activate mitotic CDK complexes and was required for mitosis (Larochelle et al., 1998). Also, the fission yeast Cdk7 homolog Mcs6/Crk1/Mop1 was isolated in a screen as a potential mitotic inducer resulting from increased Cdc2 activity (Molz et al., 1989; Molz and Beach, 1993). Analysis of a Cak1p mutant from budding yeast showed that 70% of all cells arrested at G1/S and that the remaining cells arrested in G2/M, indicating that this CAK clearly was required for both G1/S and G2/M transitions (Thuret et al., 1996; Chun and Goebl, 1997).

In plants, two CAKs have been described to date: R2 from rice and cak1At from Arabidopsis (Hata, 1991; Sauter, 1997; Umeda et al., 1998). Whereas rice R2 was shown to phosphorylate CDKs as well as the CTD of the RNA polymerase II large subunit (Yamaguchi et al., 1998), Arabidopsis cak1At displayed CAK but not CTD kinase activity in vitro (Umeda et al., 1998). Both plant CAKs are related closest to Cdk7 (Sauter, 1997; Umeda et al., 1998). In addition, they have distinct sequence stretches not found in Cdk7 or other CAKs and not shared by each other. For example, R2 has a unique C-terminal extension of ∼80 amino acids (Hata, 1991; Sauter, 1997; Umeda et al., 1998). These plant-specific CAK domains may play roles in plant-specific mechanisms of cell cycle regulation and/or DNA metabolism.

Functional analysis of the rice CAK R2 has been performed previously through yeast complementation assays. Monomeric R2 was able to rescue a temperature-sensitive Cak1p mutant from budding yeast but not a mutant form of Mcs6/Crk1/Mop1, the CAK from fission yeast (Yamaguchi et al., 1998), even though R2 is related more closely to the Cdk7-type Mcs6/Crk1/Mop1 than to Cak1p (Umeda et al., 1998).

In this study, we set out to functionally characterize the rice CAK homologous kinase R2. It was shown previously that the R2 gene is expressed at higher levels in S-phase (Sauter, 1997). We report here that R2 is regulated not only at the transcript level but also at the protein and enzyme activity levels. Furthermore, transgenic approaches were used to study subcellular localization and the effect of increased R2 on cell cycle progression in suspension cells. Increased activity in S-phase, nuclear localization, and acceleration of S-phase in R2-overexpressing cells supports the conclusion that rice R2 regulates DNA replication.

RESULTS

R2 Is a Nuclear Protein

The rice CAK R2 possesses a seven–amino acid stretch at positions 340 to 346 (PRKLRRV) that is recognized as a nuclear localization signal (NLS) by PSORT, a program designed to recognize subcellular targeting signal sequences. We used transient expression assays of a fusion protein with β-glucuronidase (GUS) as a marker protein to test the functionality of this NLS. Three constructs were made (Figures 1A to 1C, right). pGUS-R2 contained the full-length R2 coding sequence fused in frame behind GUS (Figure 1A). The second construct, pGUS-R2NLS, contained a 146-nucleo-tide fragment of R2 from nucleotides 1165 to 1311 corresponding to a 49–amino acid peptide that included the presumptive NLS fused behind GUS (Figure 1B). Finally, pGUS-R2ΔNLS contained a deletion construct of R2 without nucleotides 1052 to 1311 (i.e., without the presumptive NLS) (Figure 1C). Transcription of all three fusion constructs was driven by the constitutive 35S promoter of Cauliflower mosaic virus. Tobacco protoplasts were transfected with the 35S-GUS-R2 constructs and analyzed with a confocal laser microscope (Figures 1A to 1C, left column). The fusion proteins were detected with an antibody raised against GUS and a secondary antibody coupled to the red fluorescent dye Texas red (Figures 1A to 1C, right column). The location of the nucleus was visualized with the green fluorescent DNA binding dye chromomycin A3 (Figures 1A to 1C, middle column). A 35S-GUS-NIa fusion protein containing the tobacco etch virus NLS fused to GUS (Carrington et al., 1991) also was transformed as a control for successful transformation, expression, and nuclear localization (data not shown).

Figure 1.

Figure 1.

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Subcellular Localization of R2.

(A) Transient expression of a GUS-R2 fusion protein in a tobacco protoplast. From left to right are shown a protoplast with two nuclei; the same protoplast stained with the green fluorescent DNA dye chromomycin A3 (DNA); the same protoplast immunodecorated with an anti-GUS antibody that was visualized with a secondary antibody coupled to the fluorescent dye Texas red (fusion protein); and a scheme of the full-length R2 sequence that was fused to GUS. The NLS is indicated in detail.

(B) Transient expression of a GUS-R2/NLS fusion protein in a tobacco protoplast. A transformed protoplast is shown in a phase contrast photograph (left), after staining with the DNA dye chromomycin A3 (DNA), and after immunostaining with an anti-GUS antibody (fusion protein) as described in (A). A scheme of the R2 partial sequence containing the NLS that was fused to GUS is shown at right.

(C) Transient expression of a GUS-R2ΔNLS fusion protein in a tobacco protoplast. A transformed protoplast is shown in a phase contrast photograph (left), after staining with the DNA dye chromomycin A3 (DNA), and after immunostaining with an anti-GUS antibody (fusion protein) as described in (A). A scheme of the R2 partial sequence without the NLS-containing region that was fused to GUS is shown at right.

Full-length R2 fused to GUS resulted in translocation of the fusion protein to the nucleus (Figure 1A). Because GUS itself is a cytoplasmic protein, translocation to the nucleus was attributed to nuclear targeting mediated by R2. Translocation of the fusion protein to the nucleus was confirmed using a polyclonal antibody raised against the C-terminal half of the R2 protein (data not shown). A short fragment of the R2 protein that included the putative NLS was sufficient to target the fusion protein to the nucleus (Figure 1B). In a third experiment, a deletion construct without the presumptive NLS resulted in cytoplasmic localization of the fusion protein (Figure 1C). We conclude from these results that R2 is targeted to the nucleus and that the presumptive NLS-containing region of R2 is necessary and sufficient for nuclear targeting.

R2 Is Expressed Preferentially in S-Phase

It was shown previously that R2 transcripts accumulate rapidly in suspension cells released from a G1/S-phase block caused by the application of hydroxyurea (Sauter, 1997). mRNA levels were increased while cells moved through S-phase and then decreased. We confirmed these results using aphidicolin as a reversible inhibitor of G1/S-phase progression (Figures 2A and 2B). Within 3 hr after release from aphidicolin, R2 transcripts increased and remained increased concomitant with an increased S-phase population (Figure 2B). When cells accumulated in G2-phase, R2 transcripts decreased and reached a minimum in the subsequent G1-phase.

Figure 2.

Figure 2.

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Expression and CTD Kinase Activity of R2 in Synchronized Suspension Cells.

(A) Rice suspension cells were blocked with aphidicolin for 18 hr and then released. Samples were analyzed flow cytometrically every 3 hr. The number of cells in S- and G2-phases are given as percentages of the total cell population.

(B) RNA gel blot analysis of R2 and Os;cycB2;2 expression in partially synchronized rice suspension cells. For comparison, nonsynchronized cells (asyn) also were analyzed. Gene-specific probes were used for hybridization. Ethidium bromide–stained rRNA is shown to indicate equal loading of the gel. The scheme indicates cell cycle phase progression as measured in (A).

(C) Protein synthesis of R2 was analyzed after in vivo labeling of partially synchronized rice suspension cells. Cells were incubated with 35S-methionine at the times indicated after release from the aphidicolin block. 35S-labeled R2 protein was immunoprecipitated with an R2-specific peptide antibody from crude extract and separated by SDS-PAGE. Labeled protein was detected autoradiographically. Cell cycle phase progression is indicated in the scheme at bottom.

(D) Protein gel blot analysis of R2 was performed on partially synchronized suspension cells as indicated in (A) with an R2-specific peptide antibody. Cell cycle phase progression is indicated in the scheme at bottom.

(E) CTD kinase activity of R2 was analyzed in partially synchronized suspension cells as indicated in (A). In aphidicolin-blocked cells (0 hr) and at the times indicated after release, protein was immunoprecipitated from cell extracts with an R2-specific peptide antibody and used for in vitro phosphorylation of GST-CTD with γ-32P-ATP. Asynchronous cells also were analyzed for comparison. A sample containing preimmune serum (pre) for immunoprecipitation was included as a control for nonspecific activity. 32P-GST-CTD was separated by SDS-PAGE, and labeled protein was detected autoradiographically. Cell cycle phase progression is indicated in the scheme at bottom.

Expression of the B-type cyclin Os;cycB2;2 was included for comparison. Os;cycB2;2 was shown previously to be expressed strongly in late G2- and M-phase (Sauter, 1997). Strong expression of Os;cycB2;2 was observed between 18 and 27 hr after release from the aphidicolin block, coincident with the peak in G2-phase cells and the subsequent decline of the G2-phase population that indicates the passage of cells through mitosis.

We raised polyclonal antibodies against a peptide that corresponded to amino acids 310 to 325 of R2. The specificity of this antibody was tested using recombinant R2 and recombinant CDKs cdc2Os-1 and cdc2Os-2 as controls, overexpressed as His tag fusion proteins in Escherichia coli (data not shown). Protein synthesis of R2 was analyzed using these peptide antibodies to immunoprecipitate R2 protein from extracts of suspension cells partially synchronized with aphidicolin (Figures 2A and 2C). The cells were incubated previously with 35S-methionine to label newly synthesized protein in vivo. The synthesis of R2 protein was low in aphidicolin-blocked cells. It increased concomitant with the S-phase population within 3 hr after release and stayed increased until at least 9 hr after release. R2 protein synthesis was low in G2-phase at 18 and 24 hr after release from the G1/S-phase arrest (Figure 2C).

Protein gel blot analysis of R2 in synchronized suspension cells indicated higher protein levels in aphidicolin-blocked cells and up to 9 hr after release, while cells were engaged mainly in DNA replication (Figure 2D). When cells progressed to G2 and subsequent cell cycle phases, R2 protein levels decreased. Our results indicate that increased R2 protein abundance results from increased R2 protein synthesis, which in turn results from induction of R2 gene expression, suggesting that transcriptional control plays a role in CAK regulation in plants.

CTD Kinase Activity of R2 Is Regulated during the Mitotic Cell Cycle

Chromatographically purified R2 was shown previously to phosphorylate rice Cdc2Os-1 and human Cdk2 as well as the CTD of RNA polymerase II in vitro (Yamaguchi et al., 1998). In our assays, using unfractionated cell extracts, we failed to detect CAK activity toward Cdc2Os-1 or HsCdk2 as substrates, possibly because the sensitivity of this assay was too low (data not shown). However, we were able to analyze the kinase activity of R2 with CTD as a substrate. CTD was obtained as a glutathione S-transferase (GST) fusion protein overexpressed in E. coli. GST by itself was not phosphorylated (Yamaguchi et al., 1998).

Phosphorylation of GST-CTD by immunoprecipitated R2 protein fluctuated in a cell cycle phase–specific manner in partially synchronized suspension cells (Figure 2E). Phosphorylation was high in aphidicolin-blocked cells and up to 9 hr after release, while cells were in S-phase. High R2 kinase activity during S-phase coincided with high R2 mRNA and protein levels. Concomitant with transcript and protein abundance, R2 activity decreased between 9 and 12 hr after release from the aphidicolin block and remained low during G2-phase. Unlike RNA and protein levels, which were low throughout the remaining time, R2 activity increased again 24 hr after release, when the synchronized cells likely went through cytokinesis and entered the subsequent G1-phase. Upregulation of R2 kinase activity between 24 and 36 hr without increased R2 expression suggests post-translational regulation of R2 activity.

R2 Overexpression Accelerates Passage through S-Phase and Growth Rate

To study the effect of increased in vivo levels of R2 on cell division, suspension cells of rice were transformed with a construct that resulted in the constitutive overexpression of R2 (Figures 3A and 3B). Six independently transformed cell lines were selected and analyzed. Each line had several copies of the transgene incorporated into the genome, as analyzed by DNA gel blot analysis (data not shown). Overexpression of R2 was analyzed at the transcript level (Figure 3A) and at the protein level (Figure 3B) and was found to be increased moderately but reproducibly at both levels in all cell lines. The fact that we did not isolate lines with higher R2 protein levels may have been caused by the deleterious effects of a strong overexpression on the cells.

Figure 3.

Figure 3.

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RNA Gel Blot and Protein Gel Blot Analysis of Wild-Type and R2-Overexpressing Rice Cells.

(A) Rice suspension cells were transformed with pUbi-R2. Overexpression of R2 was analyzed in three wild-type (wt) samples (a, b, and c) and in six independently transformed cell lines (1 to 6) at 7 days after subculture using a gene-specific probe. Ethidium bromide–stained rRNA is included as a control for loading of the gel. Similar results were obtained when cells were analyzed at 5 or 12 days after subculture (data not shown).

(B) Overexpression of R2 was analyzed in protein extracts that were obtained from wild-type rice suspension cells (wt) and from six independently transformed cell lines (1 to 6). Protein was separated by SDS-PAGE, blotted, and probed with an anti-R2 peptide antibody.

We used the R2-overexpressing cell lines to study the effects of R2 overexpression on the cell division cycle. The percentage of cells in G2-phase was measured at 5, 7, and 12 days after subculture in six different cell lines (Figure 4). At all times, the average G2-phase populations were increased significantly (25 to 50%) in R2-overexpressing cells compared with those in the wild-type cells. A more detailed analysis of one transformed cell line indicated several differences with respect to cell cycle progression between transformed and nontransformed cells (Figure 5).

Figure 4.

Figure 4.

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Effect of R2 Overexpression on Rice Suspension Cells.

Rice suspension cells transformed with pUbi-R2 were analyzed flow cytometrically at 5, 7, and 12 days after subculture. Three wild-type (wt) cell lines and six independently transformed R2-overexpressing (transgenic) cell lines were analyzed. The number of cells in G2-phase was calculated as a percentage of the total cell population. Numbers given are averages (±se) from all lines analyzed in two independent experiments.

Figure 5.

Figure 5.

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Cell Cycle Phase Progression in Partially Synchronized Wild-Type Cells and in the R2-Overexpressing Transgenic Line 6.

(A) Suspension cells were partially synchronized 3 days after subculture. Aphidicolin was applied two times at 20 mg/L for a total of 18 hr to block cells at or near G1/S. Samples were taken from blocked cells (0 hr) and at the times indicated after release. Nonsynchronized cells (asyn) were used as controls. Histograms obtained with flow cytometry indicate cells with a 2C (first peak), 4C (second peak), or intermediate (between the two peaks) DNA content corresponding to G1-, G2-, and S-phase cells, respectively. The top row shows histograms from asynchronous and aphidicolin-synchronized wild-type (wt) cells up to 9 hr after release from the block; the bottom row shows transgenic line 6 treated and analyzed in the same way.

(B) Number of cells in G2-phase of the cell cycle from wild-type cells (wt; open circles) and transgenic line 6 (closed triangles) partially synchronized with aphidicolin as described in (A).

R2-overexpressing cells entered S-phase more rapidly and proceeded further into S-phase even in the presence of aphidicolin than did untransformed cells (Figure 5A). Aphidicolin is known to slow elongation at replication forks by inhibiting DNA polymerases α and δ (Levenson and Hamlin, 1993), but it does not prevent initiation at origins of replication (Mosca et al., 1992; Levenson and Hamlin, 1993). Compared with untransformed cells, which were arrested at the G1/S-phase boundary by aphidicolin, R2-overexpressing cells had begun to replicate their DNA, as indicated by flow cytometric analysis (Figure 5A).

Progression into S-phase was comparable in transformed cells between 0 and 1 hr after release and in wild-type cells at 3 hr after release (Figure 5A). Six hours after release from the aphidicolin block, all synchronized cells of the transformed line had entered G2-phase (Figures 5A and 5B). In the wild-type line, it took up to 21 hr until all cells that were released from the block had replicated their DNA and maximal G2-phase cell numbers were detected (Figure 5B), indicating that wild-type cells took much longer to replicate their DNA than did transformed cells. The overall degree of synchronization of wild-type and R2-overexpressing cells was the same. In both lines, ∼40% of all cells ultimately were detected in G2-phase (Figure 5B). The duration of G2-phase in synchronized cells was very similar in R2-overexpressing and wild-type cells (∼20 and 21 hr, respectively; Figure 5B), indicating that the increased G2-phase population in an asynchronous cell suspension did not result from a prolonged G2 period.

R2-overexpressing cells further showed a higher growth rate than did wild-type cells (Figure 6). Starting from day 4 after subculture, the transgenic line showed a significantly higher fresh weight than did the wild-type population. The difference was ∼12% at day 4 and at all subsequent time points, with the exception of the 8-day time point, at which the difference was 18%. Fresh weight increase can result from either increased cell numbers or increased cell size or a combination of both. Because cells in these rice cultures grow as aggregates, the two parameters cannot be analyzed easily. Therefore, the question of whether R2 overexpression affects cell size or cell division rate has not been resolved.

Figure 6.

Figure 6.

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Fresh Weight Increase in Wild-Type Cells and in the R2-Overexpressing Transgenic Line 6.

A total of 0.7 g each of wild-type (wt) and R2-overexpressing suspension cells was subcultured in 10 mL of medium. Total fresh weight of the two populations was determined at the times indicated up to 12 days after subculture. Results given for each time point are averages (±se) of three independent experiments with three independent samples each.

Regulation of R2 in Planta

The expression of R2 was analyzed in the intercalary meristem, in the elongation zone, and in the differentiation zone of the rice internode and compared with the expression of Os;cycH;1 that was published previously (Yamaguchi et al., 2000). Deepwater rice plants were analyzed either during normal growth (0 hr) or after induction of rapid growth by partial submergence. Transcript levels of R2 were comparable in the meristem and in the differentiation zone and were twofold higher in the elongation zone of uninduced plants (Figures 7A to 7C, 0 hr), indicating that R2 expression was not meristem specific. Expression of Os;cycH;1, on the other hand, was specific to the meristematic zone and was low in the elongation zone and in differentiated cells (Yamaguchi et al., 2000) (Figures 7A to 7C). Growth induction resulted in increased R2 and Os;cycH;1 transcript levels in meristematic but not in elongating or differentiated cells (Figures 7A to 7C). Os;cycH;1 and R2 were induced in a similar manner in the meristem 6 hr after growth induction by partial submergence (Yamaguchi et al., 2000) (Figure 7A). Induction of R2 gene expression also was observed in adventitious root initials of deepwater rice, which were induced to grow rapidly with submergence (Lorbiecke and Sauter, 1999). Thus, R2 regulation occurs in roots as well as in stems.

Figure 7.

Figure 7.

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R2 Gene and Protein Expression in the Internodes of Partially Submerged Deepwater Rice Plants.

(A) R2 and Os;cycH;1 gene expression in the intercalary meristem of deepwater rice plants that were induced to grow rapidly with partial submergence. Indicated within the graph is the progression of the growth-induced meristematic cells through the cell cycle as determined by Lorbiecke and Sauter (1998). Relative RNA abundances were calculated from RNA gel blots (Yamaguchi et al., 2000). Transcript levels shown in (A) to (C) were determined on the same RNA gel blot and therefore are comparable. Transcript levels detected in the meristem at 0 hr were set arbitrarily to 1 for each gene.

(B) R2 and Os;cycH;1 gene expression in the elongation zone of deepwater rice plants that were induced to grow rapidly with partial submergence. Transcript levels were normalized to the 0-hr time point in the intercalary meristem.

(C) R2 and Os;cycH;1 gene expression in the differentiation zone of deepwater rice plants that were induced to grow rapidly with partial submergence. Transcript levels were normalized to the 0-hr time point in the intercalary meristem.

(D) R2 protein expression in the intercalary meristem of deepwater rice plants that were induced to grow rapidly with partial submergence. Protein levels are comparable to those determined in the elongation and differentiation zones.

(E) R2 protein expression in the elongation zone of deepwater rice plants that were induced to grow rapidly with partial submergence. Protein levels are comparable to those detected in the meristem and in the differentiation zone.

(F) R2 protein expression in the differentiation zone of deepwater rice plants that were induced to grow rapidly with partial submergence. Protein levels are comparable to those detected in the meristem and in the elongation zone.

R2 protein abundance was analyzed by protein gel blot analysis in the same internodal tissues of partially submerged rice plants that were used for gene expression studies (Figures 7D to 7F). Because the kinetics of growth induction by submergence treatment are highly reproducible, the data presented in Figures 7A to 7C are comparable to those presented in Figures 7D to 7F. The tissue-specific distribution of R2 transcripts with slightly increased levels in elongating compared with meristematic or differentiated cells also was seen at the protein level (Figures 7A to 7F, 0 hr). On the other hand, R2 gene but not R2 protein expression in the meristem was induced with growth-promoting submergence treatment (Figures 7A and 7D). This may be indicative of independent regulatory mechanisms at the transcriptional and translational levels. Likewise, minor changes in protein abundance may not be detected on a protein gel blot. Constitutive R2 gene expression in the elongation and differentiation zones (Figures 7B and 7C) was accompanied by constitutive or slightly decreased protein abundance in these tissues (Figures 7E and 7F).

A second immunoreactive band with an apparent molecular mass of 34 kD was detected on protein gel blots (Figures 7D to 7F). Abundance of the 34-kD polypeptide was con-stitutive in all three tissue zones of the internode. Cross-hybridization also was observed occasionally in extracts from suspension cells with no obvious correlation to R2 levels. A 0.8-kb transcript (Hata, 1991; Sauter, 1997) that is observed occasionally in addition to the 2.1-kb R2 transcript in RNA gel blot analysis is too small to account for this 34-kD polypeptide.

We analyzed the kinase activity of R2 with GST-CTD as a substrate in the internodes of rice plants. CTD kinase activity was highest in meristematic cells and lowest in differen-tiated cells (Figure 8A). It was induced within 6 hr after submergence of rice plants in the meristem, at the time when cells entered S-phase (Figures 8A to 8C). Kinase activity in the meristem was analyzed in four independent experiments, which are summarized in Figure 8B. The average numbers (±se) indicated a significant twofold induction between 3 and 6 hr. Kinase activity remained increased thereafter. The number of S-phase cells increased between 4 and 6 hr after submergence (Figure 8C) (Lorbiecke and Sauter, 1998). In contrast, no induction of CTD kinase activity of R2 was measured in the elongation zone or the differentiation zone of the internode (Figure 8A).

Figure 8.

Figure 8.

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CTD Kinase Activity of R2 in the Internodes of Partially Submerged Deepwater Rice Plants.

(A) Autoradiographs of GST-CTD 32P-phosphorylated by immunoprecipitated R2 kinase. Kinase activity was analyzed in the intercalary meristem (IM), in the elongation zone (EZ), and in the differentiation zone (DZ) of the internodes of rice plants partially submerged for the times indicated. Relative signal intensities between the three tissues are comparable. Nonspecific background activity was determined using preimmune serum (pre) for immunoprecipitation.

(B) GST-CTD kinase activity of R2 was analyzed in the intercalary meristem (IM) of partially submerged rice plants in four independent experiments. Results were quantified densitometrically, and averages (±se) were calculated.

(C) Cell cycle phase progression of meristematic cells (IM) induced to grow rapidly by partial submergence of plants. The results were obtained by flow cytometry and have been described previously (Lorbiecke and Sauter, 1998).

R2 gene expression, Os;cycH;1 gene expression, and CTD kinase activity were induced in a similar manner in meristematic cells with growth-promoting submergence treatment (Figures 7A and 8A). Induction by submergence did not occur in the elongation and differentiation zones of the internode (Figures 7B, 7C, 7E, 7F, and 8A). However, R2 gene and protein expression were high but R2 kinase activity was low in elongating cells compared with meristematic cells, indicating some degree of post-translational regulation of R2 in elongating cells (Figures 7B, 7E, and 8A). Our analysis of rice plants, therefore, suggests that regulation of R2 activity in planta occurs at the transcriptional and protein levels.

DISCUSSION

Plants have their own lifestyle, with continued reiterative development driven by meristems. Plants cannot escape from unfavorable conditions except through growth and adaptation. Meristematic activity, therefore, is controlled by developmental signals as well as environmental conditions. Even though the basic components of the machinery that controls cell division are conserved among eukaryotes, regulation of these components has been adapted during evolution to meet the special requirements of each group of organisms. For plants, with their adaptive growth habits, we might expect greater flexibility of the cell cycle machinery and hence a greater degree of regulation of its key components, the CDKs, than what is found in animal or yeast cells.

Phosphorylation is one level of CDK regulation. CAKs are essential proteins in animals and yeast (Kaldis, 1999). CDKs have a domain, the T-loop, that blocks substrate binding (DeBondt et al., 1993). Phosphorylation of a threonine near the T-loop of CDKs by a CAK causes a conformational change that allows the substrate to bind (Russo et al., 1996). Both the T-loop and the threonine residue that can be phosphorylated are conserved in plant CDKs; therefore, it is assumed that phosphorylation catalyzed by CAKs is a general requirement for full activation of CDKs not only in animals and yeast but also in plants (Morgan, 1995).

In rice plants, the CTD kinase activity of R2 was increased in cells induced to divide rapidly but not in cells induced to elongate at a higher rate, linking control of R2 kinase activity to cell proliferation in planta. During the mitotic cell cycle, R2 levels varied at the transcript, protein, and CTD kinase activity levels. This feature is unique to plants because CAK abundance and activity are constitutive in animals and yeast. R2 was expressed and active at higher levels during S-phase, a regulation that suggests a specific role for R2 in S-phase entry and/or progression through DNA replication.

When synchronized suspension cells passed mitosis and became asynchronous, RNA and protein levels decreased, whereas CTD kinase activity recovered. Because the same antibodies were used for protein detection in protein gel blot analysis and for immunoprecipitation of kinase activity, the data suggest that CTD kinase activity is regulated post-translationally, resulting in increased activity at low apparent protein levels. In partially submerged rice plants, cells in the meristem had similar protein levels but higher CTD kinase activity than did cells in the elongation zone, strengthening the view that R2 might be subject to post-translational regulation.

A role for R2 in regulating DNA replication was supported by the observation that overexpression coincided with the accelerated passage of cells through DNA replication. These results indicated that R2 was a limiting factor for S-phase entry and/or progression in wild-type rice cells. Furthermore, R2 overexpression not only affected cell cycle progression, but it also resulted in increased growth of the cell suspension. It is not clear yet whether increased growth of the cell population originated from enhanced cell division activity or from enhanced cell growth. Premature entry into and accelerated passage through S-phase, as observed in R2-overexpressing cells, might be indicative of an accelerated cell division rate. Together, these data indicate that cell cycle progression and growth in rice are regulated through R2.

Support for a general role of CAKs in regulating DNA metabolism comes from the observation that human Cdk7 protein and Cdk7 activity are found exclusively in the cell nucleus (Darbon et al., 1994). Our analysis showed that R2 is a nuclear protein as well. Functional analysis of R2 and of its close homolog Cdk7, however, is complicated by the fact that they phosphorylate not only CDKs but also the CTD of the large subunit of RNA polymerase II. Phosphorylation of RNA polymerase II has been shown to affect transcription efficiency (Young, 1991; Carlson, 1997; Steinmetz, 1997; Leatherwood, 1998). It is conceivable that RNA polymerase II phosphorylation results in transcriptional regulation of genes required for S-phase progression (Orphanides et al., 1996).

Previous studies showed that heterologously expressed monomeric R2 protein and R2/cyclin H dimers were capable of using both CTD and CDKs as substrates for phosphorylation in vitro (Yamaguchi et al., 2000). However, the question of whether CTD or CAK kinase activity of R2, or both kinase activities, is responsible for the observed acceleration of S-phase is not resolved.

If CAK activity of R2 were involved in S-phase regulation, what would be the in vivo CDK substrates? Three CDKs have been described from rice to date (Hashimoto et al., 1992; Umeda et al., 1999). Of these, R2 phosphorylates monomeric Cdc2Os-1 but not Cdc2Os-2 or Cdc2Os-3 in vitro (Yamaguchi et al., 1998). The closest homologs of Cdc2Os-1 are Cdk1, the M-phase–promoting CDK, and Cdk2, which has functions during entry into and passage through S-phase (Mironov et al., 1999). Both Cdk1 and Cdk2 were identified as substrates of animal CAKs. Expression of Cdc2Os-1 was found to be constitutive throughout the cell cycle, indicating that it is available as a substrate at all times (Sauter, 1997; Umeda et al., 1999). Cdc2Os-2 transcript levels were abundant in S-phase, rendering it a putative in vivo substrate for R2 during DNA replication (Sauter, 1997; Umeda et al., 1999). Cdc2Os-3, on the other hand, is a G2/M-phase–specific CDK (Mironov et al., 1999; Umeda et al., 1999; Fabian et al., 2000); therefore, it is an unlikely candidate for R2-mediated S-phase acceleration.

Because R2 is a nuclear protein, its substrates also should be found in the nucleus. This is true for RNA polymerase II. For rice CDKs, subcellular localization has not been described. S-phase acceleration could result from early firing of replication origins or from an increased rate of DNA replication (Young, 1991; Orphanides et al., 1996; Carlson, 1997; Steinmetz, 1997; Leatherwood, 1998). Finally, one should keep in mind that although S-phase acceleration may be the most obvious effect of R2 overexpression, it may not be the only one.

METHODS

Plant Material

Seed of rice (Oryza sativa cv Pin Gaew 56) were obtained originally from the International Rice Research Institute (Los Baños, Philippines). Rice plants were grown as described (Sauter, 1997). For growth induction, plants were submerged in a 600-liter plastic tank filled with tap water as described, leaving ∼30 cm of the leaf tips above the water surface (Lorbiecke and Sauter, 1998). At the times indicated in Figures 7 and 8, the meristematic tissue was isolated from 0 to 5 mm above the second highest node. Tissue from the elongating zone was harvested between 5 and 15 mm above the second highest node, and differentiated tissue was harvested from the oldest portion of the internode just below the youngest node. Tissue was frozen in liquid nitrogen and stored at −70°C until use.

Fresh Weight Measurement and Synchronization of Suspension Cells

Suspension cells of the Indica rice cv IR43 were provided by G.C.G. Biswas and I. Potrykus (Swiss Federal Institute of Technology, Zurich, Switzerland) and were obtained and cultured as described (Biswas et al., 1994). The cells were subcultured weekly (Sauter, 1997).

For fresh weight measurements, 0.7 g of cells were transferred to 10 mL of medium, and total fresh weight was determined at the times indicated in Figure 6 up to 12 days after subculture. To determine the fresh weight, the medium was removed with a pipette and the weight of the cells was measured with a scale.

Suspension cells were synchronized with aphidicolin as described (Sauter, 1997; Fabian et al., 2000), with modifications. Aphidicolin is a competitive inhibitor of deoxynucleotide triphosphate binding and acts to slow DNA polymerases α and δ (Wang, 1991; Levenson and Hamlin, 1993). Three days after subculture, cells were treated with 20 μg/mL aphidicolin. After 9 hr, another 20 μg/mL aphidicolin was added to the cells. After 18 hr in aphidicolin, cells were washed three times in culture medium without inhibitor and resuspended in an equal volume of medium. At the times indicated in Figures 2 and 5, cells were harvested and either used for protoplast isolation and subsequent flow cytometric analysis or frozen immediately in liquid nitrogen for protein or RNA isolation. For in vivo protein labeling, cells were incubated for 3 hr with 75 μCi/mL (1000 Ci/mmol) l-35S-methionine (Amersham-Pharmacia, Freiburg, Germany) before harvesting and protein isolation.

Flow Cytometric Analysis

Protoplasts were obtained by treatment of suspension cells with 1% cellulase ‘Onozuka RS’ and 0.075% pectolyase Y23 (Sauter, 1997). The protoplasts were stained with 4′,6-diamidino-2-phenylindole and analyzed in a CAII cell analyzer (Partec, Münster, Germany). The relative DNA content of at least 10,000 nuclei was measured at each time point. The distribution of cells in G1-, S-, and G2-phases is shown in histograms, or alternatively, the number of cells in S- or G2-phases was calculated as percentages of the total cell population.

In Vivo Labeling of R2

Suspension cells that had been labeled with 75 μCi/mL l-35S-methionine (1000 Ci/mmol; Amersham-Pharmacia) for 3 hr were ground with sea sand in protein extraction buffer as described (Kadowaki et al., 1988). The extract was centrifuged at 15,000g for 10 min at 4°C, and the supernatant was used for immunoprecipitation.

Approximately 300 to 400 μL of supernatant was mixed with 500 μL of precipitation buffer (2% Triton X-100, 1% Nonidet P-40, 300 mM NaCl, 20 mM Tris-HCl, pH 7.4, 2 mM EDTA, pH 8, 2 mM EGTA, pH 8, 0.4 mM Na3VO4, and 0.4 mM phenylmethylsulfonyl fluoride). Polyclonal antibodies were raised in rabbit against a synthetic peptide that corresponded to amino acids 310 to 325 of R2 (Biogenes, Berlin, Germany) and used for immunoprecipitation assays and for protein gel blot analysis of R2. Five microliters of anti-R2 peptide antiserum and water was added to give a final volume of 1 mL. The protein-antibody mixture was rotated for 1 hr at 4°C. Fifty microliters of a 50% protein A–Sepharose suspension (Amersham-Pharmacia) was added and incubated for 30 min at 4°C. The suspension was centrifuged for 4 min at 4°C and 12,000g in an Eppendorf centrifuge. The pellet was washed three times with half-strength precipitation buffer. The samples were incubated for 5 min at 95°C and centrifuged for 5 min at maximal speed, and the supernatants were run on 12% SDS-PAGE. After separation, proteins were fixed in 25% isopropanol and 10% acetic acid for 30 min and incubated in amplifier solution for 30 min (Amersham-Pharmacia). The gel was dried with a gel dryer (Biometra, Göttingen, Germany) onto gel-blotting paper, covered with Saran Wrap, and exposed to x-ray film.

Kinase Assays

Crude extract was isolated from suspension cells or from the meristem, the elongation zone, or the differentiation zone of the rice internode at the times indicated in Figures 8A, 8B, and 2E. For each assay, 100 μg of protein was incubated with 10 μL of anti-R2 peptide antiserum at 4°C on a rotator (Umeda et al., 1998). As a control, 100 μg of protein was incubated with 10 μL of preimmune serum. Fifty microliters of a 50% protein A–Sepharose suspension (Amersham-Pharmacia) was added, and the mixture was incubated for another 30 min at 4°C. Protein A–Sepharose-bound antibody-protein complexes were pelleted at 700g for 5 min and washed three times with 50 mM Tris-HCl, pH 7.5, 5 mM NaF, 250 mM NaCl, 0.1% Nonidet P-40, 0.1 mM Na3VO4, 5 mM EDTA, and 5 mM EGTA. After the final wash, the pellet was resuspended in kinase buffer (25 mM Hepes-NaOH, pH 7.5, and 19 mM Mg acetate).

Kinase activity was assayed in a total volume of 30 μL essentially as described (Umeda et al., 1998). Samples in kinase buffer were incubated with 30 μM ATP, 1 μCi of γ-32P-ATP (3000 Ci/mmol), and glutathione S-transferase–C-terminal domain (GST-CTD) as substrate for 45 min at 23°C. Phosphorylation was stopped by adding SDS sample buffer and heating at 95°C for 5 min. Samples were separated by SDS-PAGE. Phosphorylation of GST-CTD was measured autoradiographically. Quantification of signals was performed with a phosphorimager using the program PCBAS 2.09g.

Protein Gel Blot Analysis

Plant material or suspension cells were frozen in liquid nitrogen and ground in extraction buffer as described (Magyar et al., 1993). The extract was centrifuged for 5 min at 4°C and 25,000g. The protein concentration in the supernatant was determined according to Bradford (1976) with the Bio-Rad protein assay. Equal amounts of protein were treated for 5 min at 95°C in 25% glycerol, 2% SDS, 25 μM DTT, 10% β-mercaptoethanol, and 0.1% bromphenol blue and separated by 12% SDS-PAGE (Laemmli, 1970).

Separated protein was transferred to a polyvinylidene difluoride membrane (Amersham-Pharmacia) for protein gel blotting or stained with Coomassie Brilliant Blue R 250 for visualization. As a control for successful transfer of the proteins, the polyvinylidene difluoride membrane was stained briefly with Ponceau red (0.2% Ponceau red, 3% trichloroacetic acid) and destained with water.

Nonspecific binding was blocked with 5% milk powder in TBST buffer (10 mM Tris, and 150 mM NaCI, 0.2% Tween-20, pH 8.0) for 1 hr at room temperature. Incubation with anti-R2 peptide antibody diluted 1:1000 was for 1 hr in TBST. Binding of the antibody was detected with a peroxidase-coupled secondary antibody at a dilution of 1:5000 in TBST and with the Renaissance Chemiluminescent Reagent Plus Kit (DuPont–New England Nuclear, Köln, Germany).

RNA and DNA Gel Blot Analysis

RNA was isolated and hybridizations were performed as described (Sauter, 1997). For RNA gel blot analysis, 25 μg of total RNA was used. The C-terminal portion of R2 or the 3′ untranslated region of Os;cycB2;2 were random prime labeled using α-32P-dCTP and used as gene-specific probes. rRNA was stained with ethidium bromide and used as a control for equal loading.

Genomic DNA was isolated from transgenic and wild-type suspension cells according to Dellaporta et al. (1983). Fifteen micrograms of genomic DNA was used for each restriction digest with SalI, SacI, or BamHI for 6 hr at 37°C. BamHI and SalI excised fragments of 1430 and 1500 bp, respectively, from the integrated DNA. SacI resulted in linearization of the integrated R2 sequence. The digested DNA was precipitated, resuspended in a small volume, separated on a 1% agarose gel, and transferred to a Hybond membrane (Amersham-Pharmacia). Hybridization was performed with the same 3′ R2-specific fragment that was used for RNA gel blot analysis. λDNA/Eco1301 Marker (MBI Fermentas, Vilnius, Lithuania) was used as a DNA size marker. DNA gel blot hybridization showed integration of several copies of the transgene in each of the six cell lines analyzed further. The hybridization patterns differed in the six lines, indicating that they were derived from independently transformed cells.

Cloning of Expression Vectors

For heterologous expression in Escherichia coli, the cDNAs of R2, cdc2Os-1, and cdc2Os-2 were cloned into pQE-32 to give His-tagged proteins (Qiagen, Hilden, Germany). All constructs were sequenced to verify in-frame ligation and correct translation of the resultant polypeptides. For use as a substrate, the CTD region of the largest subunit of RNA polymerase II from Arabidopsis (Nawrath et al., 1990) was fused to GST as described by Umeda et al. (1998). E. coli cells were transformed, and overexpression was induced with isopropylthio-β-galactoside according to instructions (Qiaexpressionist Kit; Qiagen). Nickel–nitrilotriacetic acid agarose was used for purification of the N-terminal His-tagged proteins. GST-CTD was purified using glutathione–Sepharose 4B.

For ectopic expression of R2 in rice suspension cells, the complete R2 cDNA was cloned into pUbi.cas (provided by D. Becker, University of Hamburg, Germany) between the maize ubiquitin promoter (Christensen et al., 1992) and the nopaline synthase terminator. Cells were cotransformed with a vector conferring neomycin/geneticin resistance through constitutive expression of the bacterial neomycin phosphotransferase gene (Barcelo et al., 1994). Transformation was performed with microprojectile bombardment. Transformed cells were selected in medium containing 100 mg/L geneticin. Single small colonies that derived from one cell each were chosen and put on solid medium with 100 mg/L geneticin for further selection. Six cell lines were obtained from single colonies and tested for the presence of R2 by DNA gel blot analysis. Based on the number of hybridizing bands, each cell line contained several integrated copies of R2. The transgenic cell lines were cultured subsequently in medium lacking geneticin. Further experiments were performed after many rounds of subculture in geneticin-free medium.

For nuclear localization studies, R2 was cloned into pRTL2-GUS/NIaΔBam (provided by J. Carrington, Washington State University, Pullman) (Carrington et al., 1991). This vector contains the constitutive 35S promoter of Cauliflower mosaic virus, which drives the expression of β-glucuronidase (GUS) fused to a second protein and the 35S polyadenylation signal (Restrepo et al., 1990). The original vector contains the tobacco etch virus nuclear localization signal (NLS) NIa fused behind GUS (Carrington et al., 1991). This construct was used as a positive control for nuclear targeting of GUS. For analysis of R2, NIa was replaced by in-frame fusions of full-length R2 (Figure 1A), an R2 peptide containing the putative NLS as described (Figure 1B), or by a deletion construct of R2 without the putative NLS sequence (Figure 1C). For the full-length and R2 deletion sequences, R2 was cloned without its start ATG codon behind GUS. In-frame error-free cloning was confirmed by sequencing the constructs.

Transformation of Tobacco Protoplasts and Immunofluorescence Microscopy

Rapidly growing tobacco suspension cells (Nicotiana tabacum var Xanthi) were used for protoplast formation (Fromm et al., 1986), which then were used for electroporation at a concentration of 1.5 × 106 per milliliter. Twenty micrograms of the expression vector and 100 μg of salmon sperm DNA were added to 800 μL of protoplasts before electroporation at 400 mV, 960 μF, and 200 Ω (GenePulsar; Bio-Rad). Protoplasts were resuspended in 5 mL of medium as described (Sanderfoot and Lazarowitz, 1995). Expression of the GUS fusion protein was controlled with a GUS assay as described (Sanderfoot et al., 1996).

For immunofluorescence analysis, protoplasts were treated after electroporation as described (Sanderfoot and Lazarowitz, 1995). Protoplasts were attached to slides, permeabilized with 0.5% Nonidet P-40 in 50 mM Pipes, pH 6.9, 5 mM MgSO4, and 1 mM EGTA for 30 min, dehydrated in methanol at −20°C for 10 min, rehydrated for 5 min at room temperature in 50 mM Pipes, pH 6.9, 5 mM MgSO4, and 1 mM EGTA, and blocked with 3% BSA and 0.02% azide in PBST (137 mM NaCI, 2.7 mM KCI, and 10 mM Na2HPO4, 1.8 M KH2PO4, pH 7.4) at room temperature. Immunolabeling occurred with an antibody specific for GUS (Molecular Probes, Eugene, OR) for 1 hr at room temperature in the dark. The slides were washed three times with PBST before application of the secondary antibody and incubation for 30 min in the dark. Again, the slides were washed three times with PBST and subsequently stained with 50 μM chromomycin A3, a green fluorescing DNA stain, in PBS.

Excess chromomycin A3 was washed off with PBST. Protoplasts were mounted in 90% glycerol, 0.1 M Na borate, pH 9.0, and 2.5 to 3% N-propylgallate, covered with a cover slip, and sealed with nail varnish. Protoplasts were visualized with an MRC-1000 krypton/argon dual-laser confocal system (Bio-Rad) linked to an Optiphot microscope (Nikon, Melville, NY).

Acknowledgments

We are grateful to Wiebke Hellmeyer and Silke Huss for technical assistance, to Drs. Jim Carrington (Washington State University, Pullman), Sabine Quast, and Dirk Becker (Universität Hamburg, Germany) for providing transformation vectors, and to Drs. Sondra Lazarowitz and Brian Ward (Cornell University, Ithaca, NY) for help with the nuclear localization studies. This work was supported by the Deutsche Forschungsgemeinschaft.

Article, publication date, and citation information can be found at www.plantcell.org/cgi/doi/10.1105/tpc.010386.

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