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American Society of Plant Biologists The Rice Cyclin-Dependent Kinase –Activating Kinase R2 Regulates S-Phase Progression
a Institut für Allgemeine Botanik, Universität Hamburg, Ohnhorststrasse 18, 22609 Hamburg, Germany 1 To whom correspondence should be addressed. E-mail msauter{at}botanik.uni-hamburg.de; fax 49-40-42816-229
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.
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
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
In plants, two CAKs have been described to date: R2 from rice and cak1At from Arabidopsis (Hata, 1991
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
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
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  -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-R2 NLS 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).
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
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 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
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) .
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  and  (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,
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
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.
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
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
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.
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 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
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
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
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
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 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
Flow Cytometric Analysis
In Vivo Labeling of R2 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
Kinase activity was assayed in a total volume of 30 µL essentially as described (Umeda et al., 1998
Protein Gel Blot Analysis 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
Genomic DNA was isolated from transgenic and wild-type suspension cells according to Dellaporta et al. (1983)
Cloning of Expression Vectors
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
For nuclear localization studies, R2 was cloned into pRTL2-GUS/NIa
Transformation of Tobacco Protoplasts and Immunofluorescence Microscopy 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).
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. Received September 5, 2001; accepted October 22, 2001.
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Abstract
INTRODUCTION
-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.
DNA/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. 
(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