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Previous Article | Next Article 
Molecular and Cellular Biology, August 2006, p. 6170-6184, Vol. 26, No. 16
0270-7306/06/$08.00+0 doi:10.1128/MCB.02182-05
Copyright © 2006, American Society for Microbiology. All Rights Reserved.
Department of Oncology and Cancer Research UK Labs, MRC Cyclotron Building, Imperial College London, Hammersmith Hospital, Du Cane Road, London W12 0NN, United Kingdom,1 King's College London, Department of Haematological Medicine, The Rayne Institute, 123 Coldharbour Lane, London SE5 9NU, United Kingdom,2 Department of Pathology, Cancer Studies Division, Birmingham University, Birmingham B15 2TT, United Kingdom,3 Department of Histopathology, Hammersmith Hospital & Imperial College London, Du Cane Road, London W12 0HS, United Kingdom,4 Biochemistry Building, Division of Cell and Molecular Biology, Faculty of Life Sciences, Imperial College, South Kensington, London SW7 2AZ, United Kingdom,5 Department of Virology, Faculty of Medicine, St. Mary's Campus, Imperial College London, Norfolk Place, London W2 1PG, United Kingdom6
Received 10 November 2005/ Returned for modification 9 December 2005/ Accepted 27 May 2006
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ABSTRACT |
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INTRODUCTION |
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Hematopoietic cell number is maintained by the balance between proliferation, differentiation, and death by apoptosis (35, 39). The perturbation of this equilibrium can lead to hematopoietic disorders, including autoimmune diseases, leukemias, and other myeloproliferative and lymphoproliferative disorders (35). Most of the hematopoietic progenitor and mature blood cells are relatively short lived, as they are constantly eliminated by apoptosis. To compensate for this high rate of cell loss, the hematopoietic system requires the production of 1011 human (or 108 mouse) cells each day. Besides this routine turnover, hematopoietic cell proliferation is also required at times of physiological stress. For example, red blood cell numbers increase under conditions of hypoxia, while granulocyte, macrophage, and lymphocyte populations expand during infections.
The cell cycle is under stringent control during hematopoietic cell development (14, 40). Restricting access into the cell cycle can help preserve the hematopoietic stem cell and progenitor populations from exhaustion. CKIs, including p21Cip1 and p27Kip1, are believed to set the stoichiometric thresholds for cell cycle entry in response to mitogenic stimuli, such as cytokines and interactions with stromal cells and the extracellular matrix. p21Cip1 and p27Kip1 have been shown to maintain the quiescence of hematopoietic stem cells and progenitor cells, respectively, thereby governing their pool sizes (4, 5). In addition, p27Kip1 has been demonstrated to negatively regulate the cell cycle entry of some of the more mature hematopoietic cells. However, recent mouse gene deletion studies show that, despite a pronounced lymphoid hyperplasia in thymus and spleen, p27Kip1–/– mice do not display overt hematopoietic proliferative abnormalities, indicating that other negative regulators must become activated to compensate for and to maintain hemopoietic cell numbers. Nevertheless, it is unlikely that the function of p27Kip1 is substituted by the structurally related Cip/Kip-family of CKIs, as p57Kip2 is not normally expressed at significant levels in most hematopoietic cells (29) and recent work with p21Cip1-null mice suggested that p21Cip1 does not have a role in controlling the proliferation of hematopoietic progenitors and more mature blood cells (4, 5).
A recent study proposed that p130 can function as a CDK2 inhibitor and can compensate for the loss of p27Kip1 in p27Kip1-null murine embryonic fibroblasts (MEFs) to inhibit CDK2 activity (8, 10). In MEFs with both p27Kip1 and p130 deleted, cyclin E-Cdk2 kinase activity cannot be inhibited by mitogen starvation in vitro. It is therefore possible that the lack of overt hematopoietic proliferative disorders in the p27Kip1–/– mice is due to functional compensation by the pRB-related p130 protein. However, there have been no data available to suggest that this compensatory mechanism occurs in vivo. To investigate this possibility, we crossbred the mice with p27Kip1 and p130 deleted to generate mice deficient for both p27Kip1 and p130 in order to study whether the proteins have overlapping roles in proliferative control. Since previous data from an analysis of p27Kip1–/– mice showed that p27Kip1 is a specific proliferative regulator for hematopoietic organs, including thymus and spleen (15, 23, 32), we therefore focused our study on the effects of this p27Kip1 and p130 double deletion in these hematopoietic organs, where p27Kip1 has a distinct antiproliferative role.
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MATERIALS AND METHODS |
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Hematopoietic cell isolation and staining.
Mice were sacrificed, and their bone marrow, spleens, and thymi were removed. From these hematopoietic organs, single-cell suspensions were prepared in RPMI 1640 medium by pipetting and then passing through a 70-µm nylon mesh. Cells were washed in phosphate-buffered saline (PBS) containing 0.2% bovine serum albumin, and an aliquot was taken for cell counting in a trypan blue (Sigma, Poole, United Kingdom) exclusion assay. Cells were pelleted by centrifugation for the preparation of whole-cell extracts. For the isolation of lineage-specific hematopoietic cells, cells were sequentially incubated with Miltenyi immunomagnetic microbeads specific for B220 (B cells), Gr1 (monocytes plus macrophages), or Ter-119 (erythroid cells). CD3-positive cells (T cells) were initially stained for 15 min with CD3-fluorescein isothiocyanate (FITC) as described above, followed by staining with anti-FITC immunomagnetic microbeads. Different lineages captured with the microbeads were isolated using the AutoMACS cell sorter (Miltenyi Biotec Ltd., Surrey, United Kingdom) in a two-column purification program specified by the manufacturer. Half of the positive fraction was stained using the appropriate antibody to test for hematopoietic lineage purity, and the other half was fixed for cell cycle analysis. In general, the purity of the lineage-specific cells obtained was >90%, as verified by flow cytometric analysis. For the quantification of lineage-specific hematopoietic cells, nucleated cells from spleens were labeled with lineage-specific antibodies (CD3, B220/CD45R, Gr-1, and Ter-119). The purified monoclonal antibodies anti-CD3
, phycoerythrin (PE)- or FITC-conjugated anti-B220, PE-conjugated Ter-119, and FITC-conjugated anti-Ly-6G (Gr-1) as well as the respective isotype controls were obtained from BD Pharmingen (Cowley, United Kingdom).
Histological studies and immunohistochemical staining. Following fixation in 10% phosphate-buffered formalin, tissue blocks were dehydrated in graded alcohol by using standard histological techniques, embedded in paraffin, and sectioned with a microtome. Sections were cut for immunohistochemistry onto Snowcoat Xtra slides (Surgipath, Peterborough, United Kingdom). The histological appearance was observed following counterstaining with hematoxylin and eosin (H&E) (Sigma, United Kingdom). Paraffin sections (5 µm) mounted on positively charged glass slides (VWR International, United Kingdom) were processed for conventional histology (H&E) and immunocytochemistry using the avidin-biotin-peroxidase complex (ABC) method (19). Immunocytochemistry for CD3 (rabbit anti-human polyclonal antibody, diluted 1:1,500; Dako, United Kingdom), CD45R/B220 (biotin-conjugated rat anti-mouse monoclonal antibody, diluted 1:1,000; BD Biosciences, United Kingdom), Ter-119 (rat anti-mouse monoclonal antibody, diluted 1:20; BD Pharmingen, United Kingdom), and Gr1 (rat anti-mouse monoclonal antibody, diluted 1:10; eBioscience, United Kingdom) was carried out after antigen retrieval by autoclaving sections at 126°C for 20 min according to the method of Banktlavi and coworkers (1). In brief, following autoclaving, the sections were immersed for 30 min in PBS containing fresh 0.3% (vol/vol) hydrogen peroxide, rinsed in PBS for another 30 min, and incubated for 10 min in normal goat (for CD3) or rabbit (for B220, Ter-119, and Gr1) serum diluted 1:20 in PBS. An incubation with primary antibodies was carried out overnight in a humid chamber at 4°C. The second layer of biotinylated goat anti-rabbit (for CD3) or anti-rat (for Ter-119 and Gr1) serum (1:200; Vector Labs Ltd., Peterborough, United Kingdom) and the third layer of ABC complex (Vector ABC reagent, diluted 1:100) were each applied for 30 min at room temperature. Thorough rinsing in PBS (three 5-min washes) was applied between the sequential steps of the procedure. After a final rinsing in PBS, the sections were developed and counterstained with hematoxylin. Microscopic images were captured using an Olympus DP70 camera (Shinjuku-ku, Tokyo, Japan). As a negative control for the immunostaining, nonimmune serum was used as a first layer.
B-cell isolation and culture. Splenic B cells were prepared by previously published methods (22). Briefly, T cells were killed by a cocktail of monoclonal antibodies against Thy-1, CD4, and CD8 plus guinea pig complement (25). Small, dense B cells were then isolated on Percoll density gradients, and erythrocytes were depleted by treatment with a lysis solution consisting of 0.15 M NH4Cl. These preparations typically consisted of >95% B lymphocytes. The cells were cultured at 105 cells per 100 µl in RPMI 1640 medium supplemented with 5% fetal calf serum, 2 mM glutamine, 100 U/ml penicillin-streptomycin, and 50 µM 2-mercaptoethanol and stimulated with either 10 µg/ml of polyclonal anti-immunoglobulin M (IgM) [affinity-purified F(ab')2 goat anti-mouse IgM (Cappel-ICN, Costa Mesa, CA)] or lipopolysaccharide (LPS) (Escherichia coli serotype O55:B5; Sigma, Poole, United Kingdom), 100 U/ml interleukin-4 (IL-4) (R&D Systems, Abingdon, United Kingdom), 10 µg/ml anti-CD40 (clone 3/23) (25), IL-4 plus anti-IgM, anti-CD40 plus anti-IgM, or IL-4 plus anti-CD40. Cells were collected at times indicated for poststimulation.
Cell cycle analysis. Cell cycle analyses were performed by combined propidium iodide (PI) (Sigma) and FITC (Sigma) staining as described previously (43). Briefly, cells were washed with PBS and fixed in 90% ethanol-10% PBS. Following fixation, cells were washed again and then incubated with 50 µg ml–1 DNase-free RNase (Sigma), 50 µg ml–1 PI, and 0.1 µg ml–1 FITC for 30 min at 37 C prior to analysis using a FACSCalibur flow cytometer (Becton Dickinson, Cowley, United Kingdom).
Western blot analysis and antibodies. Western blot whole-cell extracts were prepared by lysing cells with four times the packed cell volume of lysis buffer (1% Nonidet P-40, 100 mM NaCl, 20 mM Tris [pH 7.4], 10 mM NaF, 1 mM sodium orthovanadate, and protease inhibitors (Complete; Roche, Welwyn Garden City, United Kingdom) on ice for 15 min. The protein yield was quantified by using a DC protein assay kit (Bio-Rad, Hemel Hempstead, United Kingdom). A total of 50 µg of lysate was separated by sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis, transferred to nitrocellulose membranes and recognized by specific antibodies. The antibodies against cyclin D2 (M-20), cyclin D3 (18B6-10), cyclin E (M-20), CDK2 (M2), CDK4 (C-22), CDK6 (C-21), p27Kip1 (C-19), p107 (C-18), and pRB (C-15) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). The anti-phospho pRB (Ser807/811) antibody was purchased from Cell Signaling Technologies (Hitchin, United Kingdom), and the anti-phospho pRB (Thr821) antibody was from Biosource International (Camarillo, CA). The anti-p130 (anti-pRB2) (K25020) monoclonal antibodies were acquired from Transduction Laboratories (Cambridge, United Kingdom). The antibodies were detected by using horseradish peroxidase-linked goat anti-mouse or anti-rabbit IgG (Dako, Ely, United Kingdom) or mouse-absorbed goat anti-rat IgG (Southern Biotechnology Associates, Inc.) and visualized by an enhanced chemiluminescence detection system (Amersham Pharmacia Biotech, Amersham, United Kingdom).
Immunoprecipitation, CDK kinase assays, and immunodepletion.
For the immunoprecipitation and CDK kinase assays, the cells collected were washed with PBS and lysed in lysis buffer containing 20 mM Tris-HCl (pH 7.9), 150 mM NaCl, 1 mM EGTA, 1 mM EDTA (pH 8.0), 1% Triton X-100, 2.5 mM sodium pyrophosphate, 1 mM sodium orthovanadate, 1 mM phenylmethylsulfonyl fluoride. Protein lysates (100 µg) were then incubated with 5 µg of anti-CDK2 (M-2) for 2 h at 4°C. At 2 h, 50% protein A-Sepharose beads (20 µl; Amersham Pharmacia Biotech) in lysis buffer were added and the mixture was incubated for an additional 2 h. The anti-CDK2 immunoprecipitates were then washed substantially and resuspended in 20 µl of kinase buffer (20 mM Tris-HCl [pH 8.0], 10 mM MgCl2, 1 mM EDTA, 1 mM dithiothreitol) supplemented with 2.5 µg of recombinant pRB protein and 10 µCi of [
-32P]ATP (3,000 Ci/mmol; Amersham Pharmacia Biotech). Reaction mixtures were incubated for 15 min at room temperature, and the phosphorylated recombinant pRB protein was resolved with 10% SDS-polyacrylamide gel electrophoresis. The gels were then dried and exposed to X-ray films. From isolated B220+ and CD3+ cells, immunoprecipitation kinase assays were performed as described previously (21), except that cells were lysed in 20 mM Tris-HCl (pH 8.0), 250 mM NaCl, 0.5% NP-40, and 1 mM EDTA. For immunodepletion experiments, two extra immunoprecipitations were performed with either the anti-p27Kip1 antibody (C-19) or the p130 antibody. The resultant supernatants were resolved on an SDS-polyacrylamide gel electrophoresis gel and analyzed by Western blotting.
[3H]thymidine incorporation assays. Cell proliferation was monitored by [3H]thymidine incorporation assays. Small dense splenic B cells were seeded into 96-well plates at 105 cells per well and cultured in 200 µl of RPMI, 0.5 µCi [3H]thymidine, and 10% fetal calf serum. [3H]thymidine was added for the final 4 h of the indicated culture times. Cells were collected using a PHD cell harvester (Cambridge Technology, Cambridge, MA), and [3H]-thymidine incorporation into DNA was quantified by scintillation counting. Each point was determined in quadruplicate, and each growth curve was performed at least three times.
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RESULTS |
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Sections from spleen and thymus were studied from four sets of mice (wild-type, p27Kip1–/–, p130–/–, and p27Kip1–/–; p130–/–) (Fig. 3). There were no significant differences in thymus upon morphological evaluation (data not shown). Compared to other groups, spleens from p27Kip1–/–; p130–/– showed expanded red pulp containing predominantly erythroid and myeloid compartments. The red pulp-to-white pulp (consisting of primarily lymphoid cells) ratio among the p27Kip1–/–; p130–/– group was more than 1.5, while among the other groups it was less than 0.75. Furthermore, the density of nucleated cells in the red pulp was four to five times higher in the p27Kip1–/–; p130–/– group than in the wild type. Among p27Kip1–/– and p130–/– mice, the cell density in the red pulp was 1.5 to 3 times that of the wild type. The expanded red pulp of the p27Kip1–/–; p130–/– mice showed trilineage hematopoiesis in the form of groups of nucleated erythroid cells, maturing myeloid precursors, and occasional megakaryocytes. Although this was best appreciated in the p27Kip1–/–; p130–/– mice, mice from other groups also had trilineage hematopoiesis, but this was of a lesser volume and proportion.
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An increase in CDK2 activity in the splenic B220+, CD3+, Ter-119+, and Gr1+ cells of p27Kip1- and p130-deficient mice. To investigate the hematopoietic cell type(s) that contributed to the increase in CDK2 activity in the spleens of the p27Kip1–/–; p130–/– mice, we purified Ter-119+, B220+, CD3+, and Gr1+ cells from the four splenic lineages and studied the CDK2 activity by Western blot analysis using anti-phospho-pRB T821 antibodies and by immunoprecipitation kinase assays. Western blot analysis demonstrated that the CDK2 activity was up-regulated in the cells of p27Kip1–/–; p130–/– mice of all four hematopoietic lineages (Ter-119+, B220+, CD3+, and Gr1+) compared with that in the cells of p27Kip1–/–, p130–/– and wild-type mice (Fig. 7A). The results also showed that, in general, key cell cycle regulators, including p130, pRB, p27Kip1, and CDK2, in these four cell lineages were expressed in a pattern similar to that of the total splenic lysates. Immunoprecipitation kinase experiments performed with the B220+, CD3+, and Tre-119+ cells showed that, in these three cell lineages, the CDK2-associated kinase activity was up-regulated in the cells of p27Kip1–/–; p130–/– mice compared with that in their p27Kip1–/–, p130–/– and wild-type counterparts (Fig. 7B). However, we failed to perform successful immunoprecipitation kinase studies with Gr1+ cells, and this was most likely due to the limited numbers of cells purified. Nevertheless, together, the Western blotting and immunoprecipitation kinase results clearly indicated that the CDK2 activity is elevated in the cells of p27Kip1–/–; p130–/– mice in all four hematopoietic cell lineages and supported the notion that p27Kip1 and p130 cooperate to repress CDK2 activity in these cells.
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p130 is the CKI binding to CDK2 in splenocytes of p27Kip1–/– mice. While p130 is the major CKI binding to CDK2 in the splenocytes of p27Kip1–/– mice, the experiments described did not address the scenario that another CKI could have a more predominant role in binding to and inhibiting CDK2 in the p27Kip1–/– splenocytes. To investigate this possibility, we performed immunodepletion experiments through serial immunoprecipitation of cell lysates with antibodies against either p27Kip1 or p130 (Fig. 9). Three rounds of immunoprecipitation with either the p27Kip1 or p130 antibodies completely eliminated CDK2 from the splenocyte lysates. The results also demonstrated that the immunodepletion of p27Kip1 from splenocyte lysates of wild-type and p130–/– mice concomitantly removed almost all of the CDK2, whereas the CDK2 complexes in the lysates of p27Kip1–/– and p27Kip1–/–; p130–/– mice were unaffected (Fig. 9A). p130 immunodepletion caused a significant reduction of CDK2 protein from the lysates of p27Kip1–/– mice but had little effect on the CDK2 proteins in lysates from wild-type, p130–/–, and double-mutant mice (Fig. 9B). Thus, p27Kip1 is the major CKI binding to CDK2 in the splenic cells of wild-type and p130–/– mice, whereas p130 is the predominant CKI associated with CDK2 in splenocytes without p27Kip1. This result is consistent with the fact that p27Kip1 and p130 compensate for each other to regulate CDK2 activity in splenic cells. It was also notable that, after the immunodepletion of p27Kip1 and p130, a significant amount of CDK2 was retained in thymus cell lysates, suggesting that only a fraction of the CDK2 complexes was associated with p130 and p27Kip1. It is also possible, though unlikely, that most of the CDK2 is not associated with a cyclin; however, without the cyclin regulatory subunit, this pool of CDK2 will be in an inactive form and will not be a target of CDK inhibitors.
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G1S-phase controls are abrogated.
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DISCUSSION |
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The lack of overt proliferative phenotypes in the major hematopoietic organs of the p27Kip1–/– mice also indicates functional redundancy, and compensation exists to prevent excess uncontrolled proliferation (15, 23, 32). Previous studies have shown that the deletion of p27Kip1 selectively induced the proliferation of hematopoietic progenitor cells in the spleen (15). Our study demonstrated that the knockout of p27Kip1 also enhanced the cell cycle entry and proliferation of more mature B cells in response to mitogenic stimulations. Previous data demonstrated that p27Kip1 deletion leads to enhanced mitogen responsiveness in T cells in vitro (20, 30). We showed that this is also true for B cells. In this report, we provided both molecular and genetic evidence for the concept that p130 functions to compensate for the absence of p27Kip1 in binding to and inhibiting CDK2 and that p130 is a bona fide CKI in vivo. Our immunoprecipitation and immunodepletion results indicate that the pRB-related protein p130 can compensate for the absence of p27Kip1 in binding to and repressing CDK2 in spleen and thymus. This is further supported by the hyperproliferative effects observed in the splenocytes of p27Kip1 and p130 double-mutant mice compared with those seen in their p27Kip1–/– single-mutant counterparts. Our results also indicate that the compensatory mechanism does not involve the classical CKIs p21Cip1 and p57Kip2 because there was no detectable amount of p21Cip1 or p57Kip2 binding to the inactive CDK2 in the hematopoietic cells of either quiescent wild-type or p27Kip1–/– mice (data not shown). Moreover, while p21Cip1 expression levels were not elevated in the cells of p27Kip1–/– mice, p57Kip2 is not expressed at significant levels. Indeed, p21Cip1 has also been reported to have a positive rather than negative role on hematopoietic cell proliferation. The enhanced cell proliferation status and hyperphosphorylation of pRB in hematopoietic cells have been shown to be associated with an increase in p21Cip1 expression level (41). Active CDK complexes containing p21Cip1 have been detected, and it has been suggested that p21Cip1 may stabilize such complexes. Therefore, p21Cip1 may have a role independent of growth arrest in hematopoietic cells (18). Both p27Kip1 and p130 have broad antiproliferative effects on a variety of cell types and tissues outside the hematopoietic system. To eliminate the possibility of the p27Kip1- and p130-dependent environment having an influence on hematopoietic development in these mice, we used B cells as a model system to test their proliferative potential in vitro. The finding that the B cells of p27Kip1–/–; p130–/– doubly deficient mice are hypersensitive to mitogenic stimulations in vitro supports the idea that the hyperproliferation of splenocytes is not solely a result of the influence of their environment in vivo.
Previous studies have shown that a unique domain within the spacer region of p130 is essential for binding to and inhibiting CDK2 (12, 44). Although the amino acid sequence of this domain is not shared with the pRB-related p107 pocket protein, p107 has also been shown to be able to associate with and repress CDK2 through another p21Cip1-like motif located at the spacer region (2, 44, 47). Another second cyclin-binding site has also mapped to the N-terminal portions of p107 and p130 (2, 44). This N-terminal domain is capable of inhibiting cyclin-cdk2 complexes. It is therefore plausible that p107 can also functionally substitute for the absence of p27Kip1 and p130 in the p27Kip1 and p130 doubly deficient cells. In agreement with this idea, our data demonstrate that p107, like p130, can also bind to CDK2 in both spleen and thymus. Interestingly, in particular in the thymus, the amount of p107 binding to CDK2 increases when p27Kip1 is absent, indicating that p107 might also functionally compensate for p27Kip1 in repressing CDK2. This compensatory mechanism can help to explain for the lack of a significant proliferative phenotype in the p130 and p27Kip1 doubly deficient thymocytes and, possibly, for the absence of even more severe disorders in the splenocytes. In the thymocytes, the expression level of p107 is high compared with that in the splenocytes, thus allowing p107 to play a more prominent role than that of p130 in inhibiting CDK2 activity. This may explain the lack of excessive thymocyte proliferation when both p130 and p27Kip1 are lost and is supported by the finding that high levels of p107 are associated with CDK2 in the thymus but not the spleen. The high intrinsic levels of CDK2 could also potentially exacerbate the situation in the thymocytes. Indeed, functional compensation between pRB family proteins has previously been documented in vivo, but it has mostly centered on their roles in repressing E2F activity (6, 7, 11, 36). These findings also suggest that the ability of p130 to compensate for the absence of p27Kip1 is tissue and cell type specific, depending on the relative intrinsic cellular abundance of p130 and p107.
It is notable that the hyperproliferative phenotype of p27Kip1 and p130 double deletion affects only splenic cells and not thymocytes. This can be explained by the fact that the majority (>95%) of cells in the thymus are of one cell type, CD4+ or CD8+ T cells, and this one cell type contributes significantly to the phenotype of the total thymic cell population. Conversely, the fact that hyperproliferative effects of p27Kip1 and p130 deletion affect multiple splenic cell types (i.e., B and T lymphocytes, erythroid cells, monocytes, and macrophages) illustrates that this compensatory mechanism that exists between p27Kip1 and p130 is common in many different hematopoietic cells. Our data therefore suggest that cell types, such as splenocytes, that intrinsically express low amounts of p107 are largely dependent on p130 for the regulation of the CDK2 activity in the absence of p27Kip1. It is also notable that the deletion of both p27Kip1 and p130 has a more significant effect on the proliferation of the myeloid populations (erythroid cells, monocytes, and macrophages) than on the lymphoid cells (B and T cells). This probably reflects that the erythroid cells, monocytes, and macrophages or their progenitors are more dependent on p27Kip1 and p130 in their cell cycle entry control compared with the splenic T and B cells, and this can be governed by the relative expression levels of different CKIs in each cell type.
The biological significance of p27Kip1 and p130 having overlapping functions is unclear. Our previous data suggested that the repression of CDK2 activity by p27Kip1 is required for the G1 arrest induced by the inhibition of the phosphoinositide 3-kinase (PI3K) pathway (10). We also showed that the reason why p27Kip1–/– MEFs still entered cell cycle arrest after treatment with the PI3K inhibitor LY294002 is due to a compensatory mechanism by which p130 functionally substitutes for p27Kip1 deficiency (10). These results indicate that both p27Kip1 and p130 are downstream effectors of the PI3K pathway. In support of this indication, we have recently demonstrated that both p27Kip1 and p130 are direct gene targets of the FOXO subfamily of transcription factors (13, 24). The inhibition of the PI3K pathway induces a cell cycle arrest mediated by p27Kip1 and p130. The fact that both p27Kip1 and p130 function downstream of the same regulatory pathway may well be important. One possible explanation is that this compensatory mechanism exists to ensure that the cell cycle regulatory function of the PI3K pathway is intact even if one effector becomes inactivated. This compensatory mechanism may provide an essential proliferation control in settings where p27Kip1 is not expressed at normal levels, as is the case in many human tumors and hyperplasia. The cooperation between these two cell cycle inhibitors is further supported by a recent study from another group of researchers who also generated mice lacking p130 and p27Kip1 and showed that p130 and p27Kip1 cooperate to control the mobilization of angiogenic progenitors from bone marrow (42).
In summary, we show that p130 and p27Kip1 cooperate to regulate hematopoietic cell proliferation. Our data also indicate that p130 is a bona fide cyclin-dependent kinase inhibitor in vivo and functions to compensate for the absence of p27Kip1 to inhibit CDK activity. Ultimately, the generation of mice deficient for p27Kip1, p130, and/or p107 will be required to understand the compensatory mechanisms that exist to control cell cycle entry and progression in hematopoietic cells in different organs in vivo.
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ACKNOWLEDGMENTS |
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Inês Soeiro is a recipient of a fellowship from Fundação para a Ciência e a Tecnologia, Portugal. Azim Mohamedali, Nicholas Lea, and Stephen Orr were supported by the Charles Wolfson Charitable Trust. Shaun Thomas's work is supported by the Leukemia Research Fund and the Charles Wolfson Charitable Trust. David Mann and Emma Child were funded by the BBSRC. Eric Lam's work is supported by Leukemia Research Fund and Cancer Research UK.
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FOOTNOTES |
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Supplemental material for this article may be found at http://mcb.asm.org/.
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)-CD40, anti-IgM, IL-4+anti-CD40, IL-4+anti-IgM, anti-IgM+anti-CD40, LPS, or medium only for 0 (black bars), 24 (gray bars), 48 (hatched bars), and 72 (white bars) h. The cells were pulsed with [3H]thymidine for 4 h and harvested, and radioactivity was quantified. The [3H]thymidine uptake was plotted against time, and the result is representative of three individual sets of experiments. Each time point is derived from a triplicate set of results (mean ± SD [error bars]; n = 3). Due to the differences in kinetics of cell cycle entry between different treatments, Student t tests were performed on the maximum [3H]thymidine uptake points between the DKO samples and the WT, p27Kip1–/–, or p130–/– samples. There is a statistically significant difference in [3H]thymidine uptake between the DKO group and the other genotypes (P < 0.01 in response to IL-4, anti-IgM, IL-4+anti-CD40, IL-4+anti-IgM, anti-IgM+anti-CD40, LPS, and medium only but not to anti-CD40). NS, not significant. *, statistically significant (P 
0.01).