C-terminal phosphorylation controls the stability and function of p27kip1
2006; Springer Nature; Volume: 25; Issue: 21 Linguagem: Inglês
10.1038/sj.emboj.7601388
ISSN1460-2075
AutoresUta Kossatz-Boehlert, Jörg Vervoorts, Irina Nickeleit, Holly Sundberg, J. Simon C. Arthur, Michael P. Manns, Nisar P. Malek,
Tópico(s)Microtubule and mitosis dynamics
ResumoArticle19 October 2006free access C-terminal phosphorylation controls the stability and function of p27kip1 Uta Kossatz Uta Kossatz Institute for Molecular Biology, Hannover Medical School, Hannover, Germany Search for more papers by this author Jörg Vervoorts Jörg Vervoorts Institute for Molecular Biology, Hannover Medical School, Hannover, Germany Institute for Biochemistry, Klinikum der RWTH, Aachen, Germany Search for more papers by this author Irina Nickeleit Irina Nickeleit Institute for Molecular Biology, Hannover Medical School, Hannover, Germany Search for more papers by this author Holly A Sundberg Holly A Sundberg Institute for Molecular Biology, Hannover Medical School, Hannover, Germany Search for more papers by this author J Simon C Arthur J Simon C Arthur MRC Protein Phosphorylation Unit, University of Dundee, Dundee, Scotland Search for more papers by this author Michael P Manns Michael P Manns Department of Gastroenterology, Hepatology and Endocrinology, Hannover Medical School, Hannover, Germany Search for more papers by this author Nisar P Malek Corresponding Author Nisar P Malek Institute for Molecular Biology, Hannover Medical School, Hannover, Germany Department of Gastroenterology, Hepatology and Endocrinology, Hannover Medical School, Hannover, Germany Search for more papers by this author Uta Kossatz Uta Kossatz Institute for Molecular Biology, Hannover Medical School, Hannover, Germany Search for more papers by this author Jörg Vervoorts Jörg Vervoorts Institute for Molecular Biology, Hannover Medical School, Hannover, Germany Institute for Biochemistry, Klinikum der RWTH, Aachen, Germany Search for more papers by this author Irina Nickeleit Irina Nickeleit Institute for Molecular Biology, Hannover Medical School, Hannover, Germany Search for more papers by this author Holly A Sundberg Holly A Sundberg Institute for Molecular Biology, Hannover Medical School, Hannover, Germany Search for more papers by this author J Simon C Arthur J Simon C Arthur MRC Protein Phosphorylation Unit, University of Dundee, Dundee, Scotland Search for more papers by this author Michael P Manns Michael P Manns Department of Gastroenterology, Hepatology and Endocrinology, Hannover Medical School, Hannover, Germany Search for more papers by this author Nisar P Malek Corresponding Author Nisar P Malek Institute for Molecular Biology, Hannover Medical School, Hannover, Germany Department of Gastroenterology, Hepatology and Endocrinology, Hannover Medical School, Hannover, Germany Search for more papers by this author Author Information Uta Kossatz1,‡, Jörg Vervoorts1,2,‡, Irina Nickeleit1,‡, Holly A Sundberg1, J Simon C Arthur3, Michael P Manns4 and Nisar P Malek 1,4 1Institute for Molecular Biology, Hannover Medical School, Hannover, Germany 2Institute for Biochemistry, Klinikum der RWTH, Aachen, Germany 3MRC Protein Phosphorylation Unit, University of Dundee, Dundee, Scotland 4Department of Gastroenterology, Hepatology and Endocrinology, Hannover Medical School, Hannover, Germany ‡These authors contributed equally to this work *Corresponding author. Institute for Molecular Biology, Hannover Medical School, Carl Neuberg Strasse 1, Lower Saxony, 30625 Hannover, Germany. Tel.: +49 511 532 4585; Fax: +49 511 532 4283; E-mail: [email protected] The EMBO Journal (2006)25:5159-5170https://doi.org/10.1038/sj.emboj.7601388 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Entry of cells into the cell division cycle requires the coordinated activation of cyclin-dependent kinases (cdks) and the deactivation of cyclin kinase inhibitors. Degradation of p27kip1 is known to be a central component of this process as it allows controlled activation of cdk2-associated kinase activity. Turnover of p27 at the G1/S transition is regulated through phosphorylation at T187 and subsequent SCFskp2-dependent ubiquitylation. However, detailed analysis of this process revealed the existence of additional pathways that regulate the abundance of the protein in early G1 and as cells exit quiescence. Here, we report on a molecular mechanism that regulates p27 stability by phosphorylation at T198. Phosphorylation of p27 at T198 prevents ubiquitin-dependent degradation of free p27. T198 phosphorylation also controls progression through the G1 phase of the cell cycle by regulating the association of p27 with cyclin–cdk complexes. Our results unveil the molecular composition of a pathway, which regulates the abundance and activity of p27kip1 during early G1. They also explain how the T187- and the T198-dependent turnover systems synergize to allow cell cycle progression in G1. Introduction The cyclin kinase inhibitor p27kip1 plays a pivotal role in the regulation of the mammalian cell cycle. High levels of p27 expression are found in quiescent cells and the protein contributes to the maintenance of the quiescent state (Coats et al, 1996; Sherr and Roberts, 1999). Upon sufficient mitogenic stimulation p27 levels drop, thus allowing the activation of cyclin E- and cyclin A-associated cyclin-dependent kinase 2 (cdk2) (Coats et al, 1996). The levels of p27 expression throughout the cell cycle are controlled by transcriptional, translational and post-translational mechanisms (Millard et al, 1997; Kolluri et al, 1999; Montagnoli et al, 1999; Medema et al, 2000; Gopfert et al, 2003). Phosphorylation-induced degradation of p27 by the skp2-dependent SCF (skp-cullin-f-box) E3 ligase has been studied in detail. Phosphorylation of p27 at a conserved c-terminal threonine (T187) creates a binding site for the F-box protein skp2, which in concert with the SCF complex allows polyubiquitylation and subsequent proteasomal degradation of p27 (Sheaff et al, 1997; Vlach et al, 1997; Carrano et al, 1999; Sutterluty et al, 1999). Mouse fibroblasts derived from mice expressing a nonphosphorylatable form of p27 (p27T187A) reaccumulate p27T187A at the G1/S transition due to impaired turnover of the mutant form (Malek et al, 2001). However, the downregulation of p27 observed after mitogenic stimulation of quiescent cells was not impaired in the T187A cells, thereby indicating that a degradation system independent of this phosphorylation site is required for the turnover of p27 in early G1. Loss of the F-box protein skp2 in the mouse leads to a severe defect in tissue homeostasis involving multiple organs including the liver, kidney, lung and testes of homozygous skp2 knockout mice (Nakayama et al, 2000). Interestingly, while several proteins have been shown to be substrates of the skp2-dependent SCF complex, loss of p27 reverts the phenotype of the skp2 knockout mouse back to normal (Nakayama et al, 2004). Moreover, loss of skp2 prevents the degradation of p27 in primary mouse fibroblasts and liver cells after mitogenic stimulation, thereby indicating that some cell types might require skp2 for the turnover of p27 in early G1 (Kossatz et al, 2004). Meanwhile, a second ubiquitin ligase for p27, named KPC1/2, was shown to regulate the stability of p27 (Kamura et al, 2004). Degradation of p27 by the KPC ubiquitin ligase requires nucleo-cytoplasmic transport (Kamura et al, 2004); however, not all degradation of p27 takes place in the cytosol (Ishida et al, 2000; Boehm et al, 2002). In fact, confining p27 to the nucleus leads to a decrease in its stability indicating that nuclear degradation systems take part in the turnover of p27 (Rodier et al, 2001). In this work, our goal was to identify post-translational modifications on p27, which contribute to the regulation of p27 stability specifically in early G1. As our initial characterization of this early degradation system pointed towards a phosphorylation-dependent system (Malek et al, 2001), we speculated that phosphorylation of p27 at a site different from T187 might induce degradation of the protein upon mitogenic stimulation of quiescent cells. In this work, we identify the c-terminal threonine T198 as a phosphorylation site, which controls several important aspects of p27 function including binding to cyclin–cdk complexes, subcellular localization and protein stability. Interestingly, cyclin–cdk binding and protein stability are intimately linked processes, which cooperate to allow cell cycle-regulated expression of the p27 protein. Results Phosphorylation at T198 controls the stability of p27kip1 We began our study with a systematic analysis of point mutations in known or predicted p27 phosphorylation sites. To determine the consequences of these changes on p27 stability, we transfected the mutant p27 proteins into HEK 293 cells and measured their steady-state levels. Figure 1A shows the expression levels of a subset of p27 phosphosite mutants. Surprisingly, changing the threonine in position 198 into alanine (T198A) significantly decreased the steady-state expression levels of the mutant p27 protein (Figure 1A). To assay p27 phosphorylation in vivo, we raised a phospho-specific antibody against phospho-T198. As part of the characterization of this antibody, we first immunoprecipitated p27 from asynchronously growing 293 cells and showed that the phospho-T198 antibody recognizes wild-type p27 (Figure 1B). Treatment of the immunoprecipitated material with alkaline phosphatase led to a complete disappearance of the phospho-T198 signal, which was maintained when phosphatase inhibitors were added to the reaction before the addition of phosphatase. To ensure that the phospho-T198 antibody recognizes only phosphorylated T198, we transfected 293 cells with wild-type and p27T198A and showed that the phospho-specific antibody is only detecting the wild-type form of the p27 protein but not the p27T198A mutant when equal amounts of p27 protein were loaded (Figure 1C). (For more information on the characterization of the antibody, refer to Supplementary Figure 1.) Figure 1.Loss of T198 phosphorylation leads to ubiquitylation and degradation of p27kip1. (A) Expression analysis of a series of p27 phosphosite mutants after transfection into HEK 293 cells. (B) After transfection into 293 cells, the immunoprecipitated protein was either treated with alkaline phosphatase or left untreated. Phosphatase inhibitors were added (lane 3) to ensure the specificity of the reaction. Expression levels of p27 and phosphorylation status at T198 were analyzed by Western blotting. (C) After transfection of 293 cells with wild-type or T198A mutant p27, expression levels of p27 and phosphorylation status at T198 were determined by Western blotting. (D) The half-life of wild-type or T198A p27 was determined after transfection into Rat1a cells. The graph shows a half logarithmic display of the band intensities as detected by a p27-specific antibody. (E) 293 cells were transfected with p27 wild-type or the T198A mutant plasmids and increasing amounts of an UbR7-expressing plasmid. (F) 293 cells were transfected with wild-type or T198A mutant p27 and treated with the proteasome inhibitor MG132 (25 μM) for 8 h. Expression levels of wild-type and mutant p27 were analyzed by Western blotting. (G) 293 cells were transfected with p27 wild-type or the T198A mutant form with or without His-tagged ubiquitin. Ubiquitylated proteins were purified on a Co-column and detected by Western blotting with a p27-specific antibody. The graph shows a quantification of the amount of ubiquitylated p27 and p27T198A protein in relation to input protein levels. (H) HeLa cells were transfected with siRNA against skp2 and expression plasmids for p27 and p27T198A. Expression levels of p27, skp2 and actin levels were compared in siRNA-treated and untreated cells. (I) HeLa cells were transfected with siRNA against KPC1 and expression plasmids for p27 and p27T198A. Expression levels of p27, p27T198A and actin levels were compared in siRNA-treated and untreated cells. Efficient knockdown of KPC1 was verified by RT–PCR in wild-type, p27T198A and vector-transfected cells. Untransfected HeLa cells were used as positive control. Download figure Download PowerPoint To determine whether the reduced expression levels of the p27T198A mutant were due to increased turnover, we measured the half-life of p27T198A in comparison to wild-type p27 and found it to be significantly shortened (Figure 1D). Next, we tested whether the increased turnover at steady-state levels was due to increased ubiquitylation by cotransfecting HEK 293 cells with a mutant ubiquitin (UbR7), which prevents ubiquitin chain elongation. As shown in Figure 1E, UbR7 lead to a dose-dependent increase in the levels of wild type and p27T198A. To exclude an indirect effect of UbR7 on the stability of p27T198A, we directly measured the amount of His-ubiquitin incorporated into wild-type and mutant p27. As shown in Figure 1G, significantly more His-ubiquitin was incorporated into p27T198A when compared with wild-type p27 after normalization for protein input levels. Furthermore, treatment of transfected cells with the proteasome inhibitor MG132 stabilized p27T198A leading to expression levels comparable to wild-type p27 (Figure 1F). To determine whether the F-box protein skp2 is involved in the turnover of T198A mutant p27, we transfected wild-type or T198A p27 into continuously proliferating HeLa cells and reduced the expression of skp2 by siRNA-mediated knockdown. As shown in Figure 1H, reduction of skp2 expression led to an increase in the levels of wild-type and T198A mutant p27. However, the relative differences between p27 wild type and T198A protein levels were maintained under conditions of skp2 knockdown, indicating that p27T198A is still degraded by the SCF skp2-dependent degradation system, but that in addition to this pathway a second skp2-independent mechanism must regulate the abundance of the nonphosphorylatable protein in proliferating cells. Similar results were observed in skp2 KO MEFs transfected with Flag-tagged versions of wild-type and T198A p27 (data not shown). Recently, a second ubiquitin ligase named KPC1 was shown to regulate the abundance of p27 in early G1 (Kamura et al, 2004). We therefore tested whether loss of KPC1 would result in stabilization of p27T198A. As shown in Figure 1I, transfection of HeLa cells with siRNA directed against KPC1 led to a complete loss of KPC expression as detected by RT–PCR without a significant change in p27T198A levels. In summary, these results point towards a previously unknown mechanism that stabilizes p27 through phosphorylation at T198, thereby preventing ubiquitylation and proteasomal turnover. p27T198A phosphorylation controls p27 levels during the G0 and G1 phase Our transient transfection assays suggested that loss of T198 phosphorylation decreases the stability of p27 in asynchronously proliferating cells. In order to analyze the pattern of p27 phosphorylation at T198 throughout the cell cycle, we synchronized human fibroblasts in G0 using a combined serum starvation, contact inhibition protocol. As expected, p27 immunofluorescent staining was detectable in almost all cells during G0 and gradually decreased as cells progressed through G1 and S phase (Figure 2A). To determine what percentage of cells that stained positive for p27 also stained positive for phospho-T198, we performed double stainings with antibodies against p27 and phospho-p27T198. Phosphorylation at T198 was strongly detectable in approximately 60% of all quiescent cells and increased further as these cells progressed into the cell cycle (Figure 2B). Nevertheless, the remaining cells stained positive for p27 but did not show significant phospho-T198 staining. This result could suggest that not all quiescent cells express the factors required to efficiently phosphorylate p27 at T198, which might reflect different arrest stages or even the heterogeneity of the cell lines used in these experiments. However, it remains possible that the level of phosphorylation at T198 in the nonstaining cells is simply below the detection limit of the phospho-antibody. Figure 2.Generation and characterization of p27T198A fibroblast cell lines. (A) Human fibroblasts were synchronized in G0 and released into the cell cycle. To ensure specificity, we used a p27T198A fibroblast cell line (see below) as a negative control. The synchronicity of the cells was verified by flow cytometry (data not shown). At the indicated time points, cells were fixed and the number of p27-positive cells was determined by immunofluorescence staining using a p27-specific antibody. The graph shows average values of three independent experiments (in each experiment, a minimum of 300 cells were counted). A representative example of p27, and phospho-T198 immunofluorescent staining in a synchronization experiment is shown in Figure 3A. (B) Human fibroblasts were synchronized in G0 and released into the cell cycle. At the indicated time points, cells were fixed and the numbers of p27 and p27T198 double-positive cells were determined by immunofluorescence double staining using a p27 and a phospho-T198-specific antibody. The graph displays the percentage of p27-positive cells which stained also positive for phospho-p27T198 derived from several independent experiments. p27/phospho-T198 double stainings are shown in Supplementary Figure 2. (C) Introduction of a 7.7 kb genomic fragment encoding the mouse p27 locus (cartoon) into p27 knockout MEFs. (D) Analysis of the mRNA expression levels of the p27 wild-type, p27T198A mutant (T198A), wild-ype MEFs (3T3 p27 WT) and p27 knockout (p27 KO) cell lines by semiquantitative RT–PCR using the GAPDH gene as an internal control. (E) The levels of p27 or p27T198A expression in the respective cell lines (two independently derived wild-type and p27T198A lines) were measured by Western blot analysis. (F) Half-life measurement of p27 in asynchronously growing wild-type and T198A mutant cells. The graph shows a quantification of p27 band intensities of a representative experiment. (G) Identical experiment as in (E) but the cells were arrested in G0. (H) Wild-type or p27T198A mutant cell lines were arrested in G0 and then released into the cell cycle. At the indicated time points, expression levels of actin, p27, cyclin A and the retinoblastoma protein were measured by Western blotting analysis. (I) The graphs show the percentage of BrdU-positive cells detected at the indicated time points after serum readdition in wild-type and T198A mutant p27-expressing cell lines. Download figure Download PowerPoint We therefore decided to test the physiological significance of p27 phosphorylation at T198 in an in vivo model. To this end, we mutated a 7.7 kb fragment derived from the genomic mouse p27 locus at position T198 (Figure 2C) and stably integrated it or the wild-type p27 fragment into p27 knockout MEFs. To ensure that our transgenic lines recapitulate the activity of the endogenous p27 promoter, we measured p27 mRNA levels by RT–PCR. As shown in Figure 2D, all cell lines (p27 wild type and p27T198A) express p27mRNA at levels identical to unmodified wild-type MEFs. However, in agreement with our transfection data, we found that p27T198A protein was expressed at significantly lower levels than wild-type p27 in the transgenic cell lines (Figure 2E). This decrease in expression levels is due to a significantly shortened half-life of p27T198A as compared to wild-type p27 (Figure 2F). Given these observations, we concluded that phosphorylation at T198 is required to stabilize p27 in proliferating cells by preventing its proteasomal turnover. Next, we arrested wild-type and T198A cell lines in G0 and measured the expression levels of wild type and p27T198 as cells progressed through the cell cycle after mitogen stimulation. As expected, wild-type p27 was strongly expressed in G0 cells (0 h time point) and was downregulated after restimulation with mitogens (Figure 2H). However in contrast to wild-type cells, arrested p27T198A cells showed greatly reduced levels of p27T198A (Figure 2H) in G0. This reduction in protein expression was again caused by rapid degradation of the mutant protein as shown by a significantly shortened half-life (Figure 2G) of the p27T198A protein. Thus, phosphorylation at T198 protects p27 against proteolytic degradation in asynchronously proliferating and in quiescent cells. From this data we also conclude that the fact that not all quiescent cells stained positive for T198 phosphorylated p27 is most likely due to the detection limit of our antibody. Unexpectedly however after cell cycle re-entry, p27T198A levels increased continuously and peaked 12–16 h after serum stimulation. At this time point, p27T198A overall levels were similar to what is normally detected in quiescent wild-type cells (Figure 2H). This pattern was reproducibly observed in different p27T198A clones and also in pools of p27T198A transgenic cell lines (data not shown). The reaccumulation of p27T198A led to an 8 h delayed entry into S-phase compared to p27 wild-type cells as measured by BrdU labeling of synchronized MEFs (Figure 2I). This delay in the passage through the G1 phase paralleled a delay in Rb phosphorylation and in the expression of the cyclin A protein, indicating that the reaccumulation of p27T198A had a significant impact on cell cycle progression (Figure 2H and I). Stability and subcellular localization of p27 are independently regulated processes T198 phosphorylation by AKT or RSK of p27 had previously been shown to promote nucleo-cytoplasmic export of the protein (Fujita et al, 2002, 2003; Ishii et al, 2004). As subcellular localization of p27 has been shown to influence its stability (Rodier et al, 2001; Kamura et al, 2004), we analyzed whether changes in the distribution of wild-type p27 and p27T198A throughout the cell cycle might have caused the observed differences in protein expression and hence cell cycle progression. To answer this question, we first determined the subcellular expression of T198-phosphorylated p27 throughout the cell cycle in synchronized human fibroblasts using the T198 phospho-specific antibody. These experiments showed that T198-phosphorylated p27 was strongly expressed in quiescent cells (see also Figure 2B) where it is predominantly localized to the nucleus, while after mitogenic stimulation as cells passed through the G1 and S phase the T198 phospho-specific staining recognized mainly cytoplasmic p27 (Figure 3A). Figure 3.Subcellular localization of p27 is regulated through phosphorylation at T198. (A) Human fibroblasts were released into the cell cycle after 72 h of serum deprivation. At the indicated time points, the subcellular localization of p27 and T198-phosphorylated p27 was determined by immunofluorescence staining. p27 knockout fibroblasts were used as a negative control for the p27 staining. p27T198A fibroblasts were used as a negative control for the phospho-p27T198 staining. (B) The graphs display the percentage of wild-type or T198A mutant cells staining positive for p27 in either the nucleus or cytoplasm after release from quiescence. The localization of p27 in these cells was determined by immunofluorescence microscopy. At least 300 stained cells were counted per time point. Results shown are representative for at least three independent experiments. (C) Proliferating or quiescent p27 wild-type and T198A mutant cell lines were treated with MG132 or DMSO for 8 h. The subcellular localization of p27 was determined by immunofluorescence staining. (D) The indicated mutants were transfected into asynchronously growing 293 cells and the expression levels of p27 were measured by Western blotting. Download figure Download PowerPoint To determine if the loss of T198 phosphorylation affects the subcellular localization of p27, we synchronized p27 wild-type or the T198A mutant cell lines in G0. Immunofluorescence staining of p27 in G0-arrested cells revealed that both p27T198A and wild-type p27 are expressed exclusively in the nucleus (Figure 3B). However in G0 arrested cells, total protein levels of p27T198A are significantly lower than those of wild-type p27 (compare 0 h time point in Figure 2H). These results indicated that during G0, wild-type and mutant p27 are both correctly localized to the nucleus but that due to its decreased stability p27T198A is expressed at greatly reduced levels. To exclude the possibility that p27T198A, which is detected in the nucleus, only represents a stable subfraction of the total p27T198A present in the cell, we treated G0 arrested p27 wild-type and p27T198A cell lines with MG132 to prevent degradation of a putatively unstable cytoplasmic pool of the protein. As shown in Figure 3C, MG132 treatment did not increase the staining of cytoplasmic p27T198A while at the same time the total number of cells that express p27T198A in the nucleus increased (data not shown). This result therefore suggests that mislocalization is not the reason for the increased turnover of the mutant form in quiescent cells. While G0 arrested cells degrade p27T198A in the nucleus, it remains possible that delayed nuclear exit of the nonphosphorylatable form in mitogen-stimulated cells contributes to its increased stability in early G1. In this model, a cytoplasmic ubiquitin ligase would turnover p27 after its export from the nucleus, which is impaired in the p27T198A cell line. Indeed when we stimulated quiescent cells with mitogens, we observed a significant delay in the export of p27T198A (Figure 3B). Within the first 12 h of stimulation, most of the mutant p27T198A remained nuclear while at the same time its protein levels increased significantly compared to wild-type p27, which exited the nucleus within the first 12 h (Figure 3B). Between 16 and 20 h after stimulation, however 80% of the p27T198A-expressing cells showed a cytoplasmic staining while p27T198A protein levels were still significantly higher than in wild-type cells (Figures 2H and 3B). Furthermore, when we treated asynchronously proliferating wild-type and p27T198A cells with MG132 to prevent proteasomal turnover, we observed a strong increase in the cytoplasmic staining of wild-type p27 while p27T198A remained predominantly nuclear (Figure 3C). These results suggested that in quiescent and proliferating cells, p27T198A is primarily degraded in the nucleus. To test whether the predominant nuclear localization in proliferating cells is a prerequisite for the degradation of p27T198A, we constructed p27 wild-type and T198A mutants which are defective in their nuclear localization signal. Even though loss of the NLS led to a predominant cytoplasmic localization of wild-type and T198A p27 (Supplementary Figure 3A), this change in subcellular localization did not lead to a stabilization of p27T198A compared with wild-type p27 (Figure 3D) nor did it stabilize ΔNLSp27T198A compared to p27T198A (see half-life measurement in Supplementary Figure 3B). We therefore conclude that while p27T198A is predominantly degraded in the nucleus, changing its subcellular localization is not influencing the turnover of the mutant form, that is, nucleo-cytoplasmic export and degradation of p27T198A are independent processes. T198 phosphorylation controls distribution of p27 into cyclin/cdk complexes At this point of our analysis, we had shown that cell lines expressing p27T198A under the control of the genomic p27 promoter show extremely low levels of the mutant protein during G0 and in asynchronously proliferating cells. However, in synchronized cells, p27T198A accumulates during the G1 phase to levels comparable to what is regularly found in quiescent wild-type cells. Given that the expression of p27T198A during G1 had a significant impact on the start of S –phase, we wondered if p27T198A might have accumulated in cyclin/cdk complexes, which resulted in impaired Rb phosphorylation and the observed delay in G1 phase progression. We therefore measured the relative amount of p27 or p27T198A, which was bound to either cdk2 or cdk4 complexes after cells exited from quiescence and progressed through the cell cycle. For this, we immunoprecipitated p27 or p27T198A at different time points after release from quiescence and measured the amount of bound cdk2 or cdk4. Figure 4A shows a quantification of the relative amount of p27 bound into cdk2- or cdk4-containing complexes from several independent experiments (for raw data, see Supplementary Figure 4A). As expected, we found increased binding of wild-type p27 into cdk2-containing complexes after the cell enters into the G1 phase (Figure 4A). Interestingly however, at the same time, namely 12–16 h after serum stimulation, a period in the G1 phase where p27T198A levels reach their maximum, we found even more p27T198A bound into cdk2 complexes than in wild-type cells. The accumulation of p27T198A in cdk2 complexes especially between the 12 and 16 h time points led to a reduction in total cdk2- and cyclin A-associated kinase activities compared to the wild-type control cells (Figure 4B). The observed delay in cell cycle progression during the G1 phase in p27T198A cells (Figure 2I) can, therefore, be explained by a delay in cdk2 kinase activation due to the increased levels of p27T198A bound into these complexes in early G1. Given the increase in total p27 levels in p27T198A cell lines during early G1, we expected to find more p27T198A bound to cdk4 complexes. Surprisingly, however, while wild-type p27 did efficiently form complexes with cdk4, no increase in p27T198A/cdk4 complex formation was seen in the p27T198A-expressing cell lines (Figure 4A) after release from quiescence. Figure 4.p27T198 phosphorylation controls binding to cyclin/cdk complexes. (A) Wild-type or T198A mutant cell lines were made quiescent by
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