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Molecular and Cellular Biology, July 2007, p. 5079-5089, Vol. 27, No. 14
0270-7306/07/$08.00+0     doi:10.1128/MCB.00029-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Hematopoiesis and Thymic Apoptosis Are Not Affected by the Loss of Cdk2{triangledown}

Cyril Berthet,1,{dagger} Maria Cecilia Rodriguez-Galan,2 Deborah L. Hodge,2 John Gooya,3,{ddagger} Véronique Pascal,2 Howard A. Young,2 Jonathan Keller,3 Remy Bosselut,4 and Philipp Kaldis1*

Mouse Cancer Genetics Program,1 Laboratory of Experimental Immunology, Center for Cancer Research, National Cancer Institute—Frederick, Bldg. 560/22-56, 1050 Boyles Street, Frederick, Maryland 21702-1201,2 Basic Research Program, Science Applications International Corporation, National Cancer Institute—Frederick, Bldg. 560, Frederick, Maryland 21702-1201,3 Laboratory of Immune Cell Biology, Center for Cancer Research, National Cancer Institute, National Institutes of Health, Bethesda, Maryland 208924

Received 5 January 2007/ Returned for modification 1 February 2007/ Accepted 29 April 2007


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ABSTRACT
 
Cell cycle regulation is essential for proper homeostasis of hematopoietic cells. Cdk2 is a major regulator of S phase entry, is activated by mitogenic cytokines, and has been suggested to be involved in antigen-induced apoptosis of T lymphocytes. The role of Cdk2 in hematopoietic cells and apoptosis in vivo has not yet been addressed. To determine whether Cdk2 plays a role in these cells, we performed multiple analyses of bone marrow cells, thymocytes, and splenocytes from Cdk2 knockout mice. We found that Cdk2 is not required in vivo to induce apoptosis in lymphocytes, a result that differs from previous pharmacological in vitro studies. Furthermore, thymocyte maturation was not affected by the lack of Cdk2. We then analyzed the hematopoietic stem cell compartment and found similar proportions of stem cells and progenitors in Cdk2–/– and wild-type animals. Knockouts of Cdk2 inhibitors (p21, p27) affect stem cell renewal, but a competitive graft experiment indicated that renewal and multilineage differentiation are normal in the absence of Cdk2. Finally, we stimulated T lymphocytes or macrophages to induce proliferation and observed normal reactivation of Cdk2–/– quiescent cells. Our results indicate that Cdk2 is not required for proliferation and differentiation of hematopoietic cells in vivo, although in vitro analyses consider Cdk2 to be a major player in proliferation and apoptosis in these cells and a potential target for therapy.


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INTRODUCTION
 
Hematopoiesis is a tightly regulated process that involves self-renewal of stem cells, expansion of the lineage-committed progenitor population, and maturation into terminal differentiated cell types (30). Once established, the hematopoietic system supplies the organism with at least 10 different lineages in a regulated manner. Cell cycle regulation is important at each step of hematopoiesis; a low proliferation rate or quiescence is necessary for maintenance and self-renewal of primitive stem cells, a high cycling rate is required for effective expansion of the progenitor populations, and withdrawal from the cell cycle is suitable and sometimes a prerequisite for various functions of terminally differentiated cells (16). Moreover, most differentiated hematopoietic cells have the ability to reenter the cell cycle upon (antigenic) stimulation. Control of the cell cycle is required throughout the life span of the organism, since the half-life of mature hematopoietic cells varies from several hours to years. This involves continuous production of differentiated cells from hematopoietic stem cells (HSC) and the surveillance of uncontrolled proliferation that could lead to tumorigenesis. To allow normal blood development and avoid cell transformation, it is postulated that the hematopoietic cell cycle is under strict control.

Central components of the mammalian cell cycle machinery, which include cyclin-dependent kinases (Cdks), cyclins, and Cdk inhibitors, are now well characterized (1, 24, 27). Various signaling pathways affect these regulatory proteins, inducing the stimulation of hematopoietic cells and shifting the balance between proliferation and differentiation. The requirement of two sequential signals to activate peripheral blood T lymphocytes is a good example of the multilevel controls involving key cell cycle proteins. Indeed, stimulation of the T-cell antigen receptor (TCR) induces synthesis of both cyclins and Cdks that are necessary for progression through the G1 phase (15). However, expression of Cdk2/cyclin complexes is insufficient to promote S phase entry, since quiescent lymphocytes express p27, an inhibitor of Cdk2 activity (15, 29). Antigen stimulation triggers also the expression of the interleukin 2 (IL-2) receptor and competence to respond to IL-2 (33). Exposure to IL-2 of T lymphocytes in the G1 phase leads to decreased p27 expression, thereby allowing Cdk2 activation and progression into S phase (15, 29). Among the different players of the cell cycle, Cdk2 was considered the most important kinase regulating S phase. Cdk2 contributes especially to the phosphorylation of the retinoblastoma protein (Rb), which plays an important role in erythropoiesis (4, 12, 13, 21, 23) and other lineages (4, 12, 13, 21, 23). A role of Cdk2 has also been reported in the apoptosis of thymocytes, and Cdk2 might be required for antigen-mediated thymic negative selection (17, 19, 20, 35). Indeed, Cdk2 activity seems to be crucial for the induction of apoptosis in response to various stimuli (dexamethasone, heat shock, irradiation, anti-CD3, phorbol 12-myristate 13-acetate [PMA], etoposide) or cognate antigen engagement but not after anti-CD95/Fas treatment (20, 35). Bax and Bcl-2 upregulate Cdk2 activity in thymocytes undergoing apoptosis (17), and this activity is required for mitochondrial permeability disruption, cytochrome c release, and, as a consequence, activation of the downstream caspases 9 and 3 (35). All of these studies have shown that pharmacological inhibition of Cdk2 prevents induced apoptosis in thymocytes (17, 19, 20, 35). Moreover, no changes in the kinase activities of Cdk1 (Cdc2) or Cdk4 were observed after induction of apoptosis, suggesting a specific role of Cdk2 in the control of thymic apoptosis (20). However, loss of Cdk2 in vivo indicated that the Cdk2 gene is not essential, except in meiotic cells (5, 31). No developmental hematopoietic defect has been reported in previous studies, and normal thymic apoptosis has been observed in Cdk2-null mice (2). To better understand the in vivo role of Cdk2 and the apparent contradiction observed in cell culture compared to animals, we decided to address further whether inactivation of Cdk2 in mice affects apoptosis, proliferation, differentiation, or activation of hematopoietic cells.

Mouse models have been generated to aid the better understanding of the in vivo interplay of Cdks and cyclins and have suggested functional redundancy among these regulatory proteins. Results obtained from multiple knockouts of mouse Cdks or cyclins, proteins involved in the G1/S transition, have demonstrated that hematopoiesis can be affected by the loss of these proteins: (i) D-type cyclins are required for the expansion of hematopoietic stem cells, and triple knockout embryos die due to heart abnormalities combined with severe anemia (22); (ii) Cdk4 and Cdk6 double knockouts are similarly compromised in maturation of different hematopoietic lineages, and severely reduced numbers of circulating red blood cells are observed (25); and (iii) Cdk2 and Cdk4 double knockouts affect proliferation of progenitors, but to a lesser extent, by reducing the number of hematopoietic cells in this mouse model (7). These results, generated by genetic means, suggest that G1 cyclin/Cdk complexes are important for hematopoietic development. Moreover, decreased levels of Cdk inhibitors, p21Cip1 or p27Kip1, affects the renewal of stem cells (10, 11, 14, 26). Therefore, Cdk2 could be associated with specific functions in the hematopoietic system that have not been uncovered by previous phenotype screenings of Cdk2 knockout mice.

In our study, we investigated the requirement for Cdk2 in thymic apoptosis, since this role has been strongly suggested by pharmacological inhibitor experiments (17, 19, 20, 35). We induced apoptosis in Cdk2-null thymocytes and compared the results with thymocytes treated with roscovitine, an inhibitor of Cdk2 that protects thymocytes from apoptosis. Using genetic or pharmacological means, we analyzed also the potential compensation by other Cdks. Our second approach was to analyze the subpopulations of the different hematopoietic compartments and to challenge these subpopulations to determine whether their activation capacity was affected by the loss of Cdk2.


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MATERIALS AND METHODS
 
Generation of Cdk2–/– mice. Heterozygous Cdk2+/– mice were backcrossed to C57BL/6 mice, which were in turn intercrossed to generate homozygous mutant Cdk2–/– animals (5). Most mice used in this study were of a mixed C57BL/6 x 129S1/SvlmJ background (N2 backcrossed), and our results were confirmed with N6 backcrossed mice (considered to be of a C57BL/6 background). Mice were routinely genotyped according to a previously described protocol (7). Mice were group housed under standard conditions with food and water available ad libitum and were maintained on a 12-h light/dark cycle. Mice were fed a standard chow diet containing 6% crude fat and were treated humanely, in compliance with National Institutes of Health guidelines for animal care and use.

Cell preparation and culture. Bone marrow cells and splenocytes were prepared from 3- to 6-month-old mice, thymocytes from 6- to 12-week-old mice. The thymus and spleen were dissociated, while bone marrow cells were flushed and cells were passed through fine mesh (catalog no. 352340; BD Falcon) to generate single-cell suspensions. The cells were counted and resuspended in phosphate-buffered saline with 0.1% bovine serum albumin (Sigma-Aldrich, St. Louis, MO) or in cell culture medium. Cell culture was performed in a humidified 5% CO2 atmosphere at 37°C in RPMI 1640 medium (21870; Invitrogen) supplemented with 10% heat-inactivated fetal calf serum, 2 mM L-glutamine (0567; Invitrogen), 100 IU/ml penicillin, and 50 µg/ml streptomycin (15140-122; Invitrogen). Thymocytes were washed and cultured in suspension for 24 h in supplemented medium with the addition of 10 µM ß-mercaptoethanol. Cells were treated as indicated with dexamethasone (1 µM) (265005; Calbiochem), etoposide (2.5 or 10 µg/ml) (33419-42-0; Sigma), PMA (10 ng/ml) (524400; Calbiochem), and/or roscovitine [2-(R)-(1-ethyl-2-hydroxyethylamino)-6-benzylamino-9-isopropyl-purine] (557364; Calbiochem); Cdk2 inhibitor III [2(bis-(hydroxyethyl)amino)-6-(4-methoxybenzylamino)-9-isopropyl-purine] (238803; Calbiochem); and Cdk1 inhibitor III {ethyl-(6-hydroxy-4-phenylbenzo[4,5]furo[2,3-b])pyridine-3-carboxylate} (217697; Calbiochem). All Cdk inhibitors were dissolved in dimethyl sulfoxide. Splenocyte suspensions were cleared of red blood cells by incubating them for 1 min in ACK lysis buffer (10-548E; BioWhittaker) and cultured in supplemented medium. Adherent cells were isolated from nonadherent cells by incubation in plastic flasks for 1 h at 37°C. Nonadherent cells were recovered by gently washing them with warm medium and were cultured in suspension for 24 to 72 h. Nonadherent cells were stimulated with anti-CD3 and anti-CD28 antibodies. To induce macrophage differentiation, bone marrow cells were cultured for 6 days in supplemented medium with the addition of 5% horse serum, 1 mM sodium pyruvate (03810; Invitrogen), and 10 ng/ml macrophage colony-stimulating factor (mCSF) (315-02; Peptech). After 6 days, cells were cultured in the presence of 100 ng/ml gamma interferon (8311; eBioscience) for 24 h and stimulated with 1 to 20 ng/ml lipopolysaccharide (LPS) (011:B4; Sigma) for another 24 h.

Cell staining and flow cytometry analysis. Cells were resuspended at a density of 1 x 106 cells/100 µl in phosphate-buffered saline with 0.1% bovine serum albumin (Sigma-Aldrich, St. Louis, MO) and stained with the indicated antibodies. Throughout staining, the cells were kept at 4°C and antibodies were used at a concentration of 0.5 µg/1 x 106 cells. Cell fluorescence was analyzed with fluorescence-activated cell sorter (FACS) LSRII or a FACSCalibur (BD Biosciences) with 4-decade logarithmic amplification. Live cells were identified by forward light scatter and propidium iodide (PI) gating. Apoptosis was detected with the caspase 3 intracellular activity assay kit I (PhiPhiLux G1D2 [235430]; Calbiochem). Approximately 5 x 105 cells were pelleted by centrifugation at 600 x g in a microcentrifuge. The supernatant was removed, and the cell pellets were gently resuspended in 50 µl of PhiPhiLux SG1D2 substrate solution. The cells were incubated in open tubes in a 5% CO2 incubator at for 1 h. The cells were washed once in ice-cold FACS buffer and resuspended in FACS buffer. PI was added, and flow cytometry analysis was performed with the FL1 channel setting to examine caspase 3 cleavage of the PhiPhiLux SG1D2 substrate.

Antibodies. The following monoclonal antibodies were from BD-Pharmingen: antibodies against TCRß (anti-TCRß; H57-597), anti-CD3 (145-2C11) anti-CD4 (RM4.4 and GK1.5), anti-CD5 (53-7.3), anti-CD8{alpha} (53-6.7), anti-CD28 (37.51), anti-CD69 (H1.2F3), anti-B220 (RA3-6B2), anti-TER119 (TER-119), anti-c-Kit (2B8), anti-GR-1 (RB6-8C5), anti-F4/80 (6F12), anti-HSA (M1/69), anti-CD25 (2A3), and anti-Fc{gamma}RII/III (2.4G2). Other monoclonal antibodies used were anti-Sca-1 (Ly-6A/E; eBioscience) and anti-Mac-1 (M1/70). All antibodies used for FACS analysis were conjugated with the appropriate fluorochrome.

CFU assay (CFU-c). Fresh bone marrow cells were plated in Iscove's modified Dulbecco's medium (Gibco-BRL), 10% heat inactivated fetal bovine serum, 100 units/ml penicillin, and 50 µg/ml streptomycin, with 0.35% (wt/vol) SeaPlaque agarose (Cambrex Bio Science Rockland, Inc. Rockland, ME). The following hematopoietic growth factors were added according to the specific assay: 100 ng/ml murine stem cell factor (mSCF), 50 ng/ml murine IL-6 (mIL-6), 30 ng/ml mIL-3, and 30 ng/ml murine granulocyte-macrophage CSF (mGM-CSF) (all growth factors were from Peprotech Inc., Rocky Hill, NJ). Cells were plated at a density of 2 x 104 cells/ml/plate in 35- by 10-mm cell culture dishes (Nunc), with a 2-mm grid to facilitate counting, and were incubated at 37°C and 5% CO2 for 12 days.

Bone marrow transplantation assay. Bone marrow cells were harvested from littermate donor mice (Ly5.1), and 5 x 105 Cdk2–/– or Cdk2+/+ cells were mixed with equal numbers of C57BL/6 bone marrow cells (competitors, Ly5.2). Cells were transplanted into irradiated (950 rad) C57BL/6 (CD45.1) mice with support marrow from CD45.1 mice. Hematopoietic reconstitution was determined by analyzing hematopoietic cells from peripheral blood, bone marrow, spleen, and thymus at 46 weeks after transplantation by flow cytometry. Briefly, cells were stained with antibodies that recognize donor (Ly5.1) or host (Ly5.2) cells in combination with phycoerythrin-conjugated antibodies, including Gr-1, B220, CD4, TER119, immunoglobulin M (IgM), CD19, and TCR{alpha}ß, or biotin-conjugated antibodies, including Mac-1, CD3{varepsilon}, CD8, CD71, IgD, and TCR{gamma}. All antibodies and tricolor streptavidin used for lineage determinations were from BD Pharmingen.

Serum cytokine quantitation by CBA. Supernatants from macrophage cell cultures were harvested, and 50 µl was analyzed for cytokines with the mouse cytometric bead array (CBA) inflammation kit (BD Biosciences, Immunocytometry Systems, Chicago, IL) on a FACScan flow cytometer affixed with a 488-nm laser (Becton Dickinson Immunocytometry Systems, Mountain View, CA), according to the manufacturer's suggested protocol.

Kinase assays and Western blotting. Proteins from thymocytes were isolated at 4°C with the following lysis buffer: 50 mM HEPES (pH 7), 150 mM NaCl, 2.5 mM EGTA, 1 mM EDTA, 10 mM ß-glycerol phosphate, 0.5% Tween 20, 10% glycerol, 1 mM dithiothreitol, 2 mM NaF, and 1x protease inhibitors (10 µg/ml each of leupeptin, chymostatin, and pepstatin [Chemicon, Temecula, CA]). Lysates were centrifuged for 45 min at 18,000 x g and 4°C, and supernatants were frozen in liquid nitrogen. Protein concentrations were determined with the Bradford protein assay (500-0006; Bio-Rad). Lysates were analyzed by immunoblotting, immunoprecipitation, and kinase assays with the substrate histone H1, as previously described (5). Affinity-purified antibodies against Cdk2, Cdk1, Cdk4, cyclin B, and p27 have been described (5). Other antibodies used, rabbit anti-cyclin A (0.2 µg/ml) (H-432 [Sc-751-AC]; Santa Cruz), are commercially available.


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RESULTS
 
Cdk2 is not required for apoptosis. The thymus is the hematopoietic compartment where maturation of T lymphocytes occurs. It requires several waves of proliferation, followed by positive and negative antigen selection. These steps of selection are essential for the immune system to develop a proper allogeneic response and avoid autoimmune reactions (8, 34). The selection leads to massive apoptosis of T lymphocytes that do not recognize the major histocompatibility complex (positive selection) or bind with high affinity to self-antigens (negative selection). It has been suggested that Cdk2 activity is required for antigen-mediated thymic negative selection (17, 19, 20, 35). Previously, we did not observe an increase of apoptosis in vivo in Cdk2–/– thymus (2); however, this observation was based only on terminal deoxynucleotidyltransferase-mediated dUTP-biotin nick end labeling. This assay might not necessarily eliminate the possibility of increased apoptosis, as apoptotic thymocytes are quickly engulfed and eliminated in vivo. Therefore, we investigated whether Cdk2-null thymocytes would be protected from apoptosis in culture.

Cdk2 activity was found to be increased after induction of apoptosis in thymocytes, and pharmacological inhibition of Cdk2 activity protects cells from death (17, 19, 20, 35). These results suggested that Cdk2 contributes to thymocyte apoptosis; to extend these previous in vitro results, we reproduced these experiments using Cdk2–/– thymocytes instead of chemical inhibition of Cdk2 activity. We monitored apoptosis through caspase 3 activity assays. Our results demonstrated that Cdk2–/– thymocytes undergo apoptosis after treatment with 1 µM dexamethasone (Fig. 1A). Cdk2 was not required for this process, since apoptosis induced by 1 µM dexamethasone was similar in Cdk2–/– and wild-type thymocytes (Fig. 1A). Moreover, we confirmed that roscovitine protects wild-type thymocytes from apoptosis, but roscovitine also prevented cell death in Cdk2–/– cells (Fig. 1A). This result suggested that roscovitine can act through pathways other than inhibition of Cdk2 activity. This inhibitor is not very specific at the concentration used (50 µM, similar to that used previously [17, 20]), and so it is possible that roscovitine inhibits other Cdks that compensate for Cdk2. To determine the pathway inhibited by roscovitine, we performed a similar experiment using Cdk4–/– thymocytes, since Cdk2 and Cdk4 have common functions (7). Cdk4–/– thymocytes behaved like Cdk2–/– and wild-type thymocytes: they undergo apoptosis after dexamethasone treatment and survive in the presence of roscovitine (Fig. 1B). Cdk4 is most likely not the main target of roscovitine; however, we would need double knockout thymocytes to be sure that Cdk2 and Cdk4 cannot compensate for each other in this pathway. Double knockout of Cdk2 and Cdk4 leads to embryonic lethality (7); therefore, it was not possible to test this hypothesis. Another potential target is Cdk1 (Cdc2), a Cdk involved in mitosis that has been shown to compensate for Cdk2 (2). To determine by genetic means whether compensation of Cdk2 by Cdk1 is responsible for the resistance of Cdk2-null thymocytes to apoptosis, Cdk1-null cells would need to be analyzed; unfortunately, they are not currently available. As an alternative approach, we performed an experiment with a Cdk1 inhibitor (Cdk1 inhibitor III [Calbiochem]; see Materials and Methods) and found no rescue of apoptosis after dexamethasone treatment in wild-type or Cdk2-null thymocytes (Fig. 1B). This inhibitor alone has no effect on the survival rate of thymocytes (data not shown). This suggests that Cdk1 does not compensate Cdk2 in this process or that the Cdk1 inhibitor does not affect the same targets as roscovitine. Several additional observations corroborate these results: (i) no change in apoptosis level was detected in the thymus of Cdk2–/– p27–/– mice (2), where Cdk1 compensates for the absence of Cdk2 activity; (ii) increased activity of Cdk2 and Cdk1 (due to the lack of p27) had no effect on apoptosis (2); and (iii) comparison of Cdk2–/– with p27–/– thymi indicates that apoptosis in vivo is not affected by the lack of Cdk2.


Figure 1
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  		FIG. 1. Apoptosis of Cdk2–/– thymocytes. (A) Apoptosis of thymocytes was analyzed by flow cytometry. Cells were stained with propidium iodide (blue), and caspase 3 activity (green and blue) was detected. Viable cells (red) are double negative. Cdk2–/– and Cdk2+/+ thymocytes were harvested and treated for 24 h with 1 µM dexamethasone (Dex) to induce apoptosis and/or 50 µM roscovitine, which inhibits Cdk2. Percentages of each population are indicated inside each gate. (B) A similar assay was reproduced in five independent experiments (n ≥ 6). Cdk4-null thymocytes were included (n = 2). Roscovitine (Rs) was added at 50 or 12.5 µM. Cdk2 inhibitor (inh) III was added at 20 µM, and Cdk1 inhibitor III was used at 7 µM. Percentages of viable nonapoptotic cells were plotted.

Finally, we aimed to determine whether a reduced concentration of roscovitine or another Cdk2 inhibitor (Cdk2 inhibitor III [Calbiochem]; see Materials and Methods) would reduce the levels of cell death. Roscovitine was used at 12.5 µM, which is considered more specific for inhibition of Cdk2 (3), instead of the commonly used concentration of 50 µM. The survival rate of dexamethasone-treated lymphocytes was similar in wild-type, Cdk2–/–, and Cdk4–/– cells but lower than in cells protected with 50 µM roscovitine compared to 12.5 µM (Fig. 1B). This observation suggested that roscovitine protects cells against apoptosis at a concentration that might not be specific for Cdk2. On the other hand, Cdk2 inhibitor III protected thymocytes from dexamethasone-induced apoptosis (Fig. 1B). Both inhibitors are purine analogs and may share similar specificity toward other kinases. Together, these results suggest that Cdk2 inhibitors affect survival of thymocytes but most likely through the inhibition of a kinase other than Cdk2. This kinase may even not be a Cdk, as Cdk4 and Cdk1 appear to not be involved in the survival rate of thymocytes after dexamethasone treatment.

FACS analysis did not determine whether there was a difference in the kinetics of how wild-type and Cdk2–/– cells die. Therefore, we performed a time course of the experiment shown in Fig. 1A and determined the viability of thymocytes (Fig. 2A). We found that the kinetics of cell death and the protection by roscovitine are similar in wild-type and Cdk2–/– cells. To verify that the Cdk2–/– thymocytes did not express the Cdk2 protein, as expected from genotyping (data not shown), we performed Western blot analyses (Fig. 2B). Cdk1, Cdk4, and p27 were expressed at identical levels in both cells, whereas Cdk2 was expressed only in wild-type cells (Fig. 2B). We then analyzed the effect of roscovitine treatment on Cdk activity (Fig. 2C). Kinase activity associated with cyclin A and Cdk2 was reduced after roscovitine treatment (Fig. 2C, top two panels; compare lanes 2 and 4), but interestingly cyclin B and Cdk1 kinase activity was unaffected. As expected, Cdk2 activity was detected in wild-type thymocytes but not in Cdk2–/– cells (Fig. 2C, second panel from top). Our results indicate that roscovitine potently blocks apoptosis but may not act through the inhibition of Cdk2 and/or Cdk1.


Figure 2
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  		FIG. 2. Molecular analysis of apoptosis in thymocytes. (A) Viability of thymocytes over a 24-h time period was determined. Wild-type untreated thymocytes (Cdk2+/+) (contr), untreated Cdk2–/– thymocytes, Cdk2+/+ thymocytes treated with dexamethasone (Dex), Cdk2–/– thymocytes treated with dexamethasone, Cdk2+/+ thymocytes treated with dexamethasone and roscovitine (Rosc), and Cdk2–/– thymocytes treated with dexamethasone and roscovitine were plotted. (B) Western blots of Cdk2+/+ and Cdk2–/– thymocyte extracts probed with antibodies against Cdk1, Cdk2, Cdk4, and p27. Molecular size markers are indicated on the left, in kilodaltons. (C) Kinase activity was determined by immunoprecipitating thymocyte extracts with the antibodies indicated on the right, followed by an in vitro histone H1 assay using radiolabeled ATP.

To extend our results, we analyzed apoptosis of T lymphocytes after antigenic stimulation. We treated splenocytes with anti-CD3 and anti-CD28 antibodies and analyzed caspase 3 activity in stimulated lymphocytes. Similar viability was observed in wild-type and Cdk2–/– splenocytes after 3 days of stimulation (Fig. 3A), suggesting that Cdk2 is not essential for apoptosis of stimulated T lymphocytes. Furthermore, we tested whether roscovitine rescues apoptosis induced by stimuli such as etoposide (which induces DNA damage) or PMA (a phorbol ester that induces proliferation) in thymocytes. As we have shown for dexamethasone (Fig. 1), roscovitine protected cells potently from cell death induced by PMA and etoposide (Fig. 3B), indicating a general mechanism.


Figure 3
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  		FIG. 3. Alternative stimuli leading to apoptosis. (A) Caspase 3 activity in splenocytes was measured at day 0 and after 3 days of in vitro stimulation with anti-CD3 and anti-CD28. Percentages of apoptotic cells were plotted. Error bars represent an average of three independent experiments (n = 8). (B) Apoptosis in Cdk2+/+ and Cdk2–/– thymocytes was induced by treatment with etoposide (Etop) (2.5 or 10 µg/ml) or PMA (10 ng/ml) in the presence or absence of 50 µM roscovitine (Rs). Cell death was determined by a FACS-based caspase 3 assay (see Materials and Methods).

Thymocyte development in Cdk2-null mice. We then analyzed the thymic subpopulations to determine if their development was impaired. Thymic maturation of T lymphocytes is correlated with the rearrangement and expression of the TCR associated with the costimulatory receptors CD4 and CD8. We analyzed the expression of these two markers during the various stages of T-cell differentiation that include the double-negative early stage, the double-positive selection stage, and the single-positive final stage of differentiation. Intermediate expression of CD4 or CD8 is also observed (CD4lo CD8hi or CD4hi CD8lo), but the decrease in cell surface expression of CD4 and CD8 molecules cannot be considered a progressive decrease of expression that assesses a lineage choice in immature thymocytes. Indeed, CD8 expression is down-regulated by both CD4 and CD8 differentiating cells, causing thymocytes that are undergoing selection into either lineage to adopt a transitional CD4hi CD8lo expression (8). We isolated thymi from 6- to 9-week-old animals and counted similar numbers of cells in Cdk2 knockout and wild-type thymi. In Cdk2-null thymi, we observed normal percentages of double-negative and double-positive cells, suggesting that the first wave of T-lymphocyte proliferation, followed by TCR rearrangement, occurs properly (Fig. 4A). Lymphocytes with intermediate profile expressions of CD4 or CD8 (CD4hi CD8lo or CD4lo CD8hi) are comparable in Cdk2–/– and wild-type cells (Fig. 4A). The initiation of terminal maturation is most likely not altered in Cdk2-null thymocytes, since it has been suggested that the CD4hi CD8lo population is an intermediate early stage occurring during the maturation of CD8 or CD4 single-positive lymphocytes (8). We also observed a similar percentage of CD8 or CD4 single-positive lymphocytes (Fig. 4A). This profile of expression suggested that differentiation of thymocytes is not impaired in vivo, although Cdk2 is ablated. To complete our study, we performed further investigations with additional specific thymic differentiation markers and analyzed marker expression for each thymic subpopulation (Fig. 4B). Expression of HSA (CD24), a marker of immature T lymphocytes that decreases during differentiation, was similar in both genotypes. It is highly expressed in double-negative cells and reduced in double-positive cells, CD4+ and CD8+ (Fig. 4B). CD5 is associated with maturation of single-positive lymphocytes, and its expression was enhanced in Cdk2–/– and Cdk2+/+ cells (Fig. 4B). CD69 is a signaling molecule that is expressed shortly after the stimulation of double-positive thymocytes, which appears to be similar in both genotypes (Fig. 4B). Populations with intermediate CD4 or CD8 expression also showed a similar pattern (data not shown). Finally, expression of H57 (mature TCR expressed at a late stage of lymphocyte maturation) was enhanced along the maturation process and was expressed in all single-positive cells (>95%) (Fig. 4B). This pattern suggested that the lack of Cdk2 does not affect a specific stage of thymocyte development and led to the differentiation of CD4+ or CD8+ cells in a range identical to that of wild-type mice. These observations also confirm that Cdk2 is not required for thymic apoptosis, since none of the subpopulations was overrepresented.


Figure 4
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  		FIG. 4. Analysis of thymocyte subpopulations in Cdk2–/– mice. (A) Analysis of CD4 and CD8 receptor expression by flow cytometry in thymocytes to discriminate subpopulations in Cdk2–/– mice (n = 9) compared to their wild-type littermates (n = 11). Dot plots of representative data are shown, and average percentages of each subpopulation acquired in three independent experiments are plotted. DN, double negative; DP, double positive. (B) Expression of specific markers (HSA, CD5, CD69, H57) in the indicated subpopulations. DN, double negative; DP, double positive.

Stem cells and progenitors in Cdk2-null bone marrow. The microenvironment of adult bone marrow supports a dual biological role: the self-renewal of HSC and the expansion of progenitors. These roles lead to opposite signals for the cell cycle, with a low rate of division for HSC and a high proliferation rate for progenitors. The INK4 family of inhibitors and p27 contribute to the maintenance of quiescence in HSC, and decreased expression levels of these inhibitors, associated with increased Cdk2 activity, are required for reentry into the cell cycle (11, 14, 26, 37). The lack of p21 in stem cells impairs their self-renewal capacity and leads to premature stem cell depletion (10). On the other hand, p21 intracellular levels increase in hematopoietic progenitors upon mitogenic cytokines, which correlates with enhanced proliferation of progenitors and release of p27 from Cdk2 (26). In these cells, proliferation is significantly reduced when p21 is lacking (26). The stoichiometric interplay of p21 and p27, bound to Cdk2, plays a critical role in the mitogenic response of progenitors cells, which is important for normal maintenance of the stem cell and progenitor cell compartments (11). We wondered whether the lack of Cdk2, the direct target of p21 and p27, would affect these hematopoietic compartments. To address this question, we analyzed several biomarkers by flow cytometry to determine the ratio of HSC and bone marrow progenitors in Cdk2-null mice to control wild-type littermate mice. The numbers of bone marrow cells were similar between both types of mice, and we did not observe any significant difference in c-kit+/Sca-1+ or c-kit+/Sca-1– cells (Fig. 5A), suggesting that in adult mice the HSC and progenitor populations, respectively, are normal in the absence of Cdk2. We then analyzed different lineages—macrophages (Mac-1+), neutrophils (Gr-1+), erythroid cells (Ter119+), and B cells (B220+)—without noticing changes in the flow cytometry profile (Fig. 5A). Apparently, Cdk2 does not affect the proportion of stem/progenitor cells, and to confirm this observation we performed CFU assays with specific cytokine cocktails, adapted to induce expansion of different hematopoietic subpopulations. We focused our study on multipotential progenitors (SCF, IL-3, and IL-6) and progenitors more committed toward macrophage (mCSF), granulocyte (SCF and G-CSF), and granulocyte-macrophage (IL-3 + GM-CSF) lineages. Similar numbers of colonies were observed for Cdk2–/– and wild-type cells in each subpopulation analyzed (Fig. 5B), suggesting that Cdk2 functions are compensated for in in vitro proliferation of primitive and committed myeloid progenitors following cytokine stimulation.


Figure 5
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  		FIG. 5. Analysis of bone marrow cells in Cdk2–/– mice. (A) Analysis of bone marrow subpopulations by flow cytometry in Cdk2–/– mice (n = 5) compared to their wild type-littermates (n = 4). Dot plots of representative data are shown, and average percentages of each subpopulation acquired in three independent experiments are plotted. (B) CFU-c assays performed with Cdk2–/– bone marrow cells (n = 5) and those from their wild-type littermates (n = 4) using the indicated cytokines. Each assay was performed in triplicate. Averages of colony numbers obtained from one experiment are plotted. Two other independent experiments yielded comparable results. (C) Competitive grafts in five recipients with Cdk2–/– (n = 2) or wild-type (n = 2) bone marrow cells were performed, and cells were analyzed after 46 weeks. Repopulation of the indicated compartments is expressed by the percentage of the Ly5.1 subpopulation. Myeloid cells and B cells correspond to the Mac-1+/Ly5.1+ and B220+/Ly5.1+ subcompartments, respectively.

To determine whether Cdk2-null HSC can expand and generate all hematopoietic lineages in vivo, we performed long-term bone marrow transplantation assays. Such in vivo assays allowed us to monitor the capacity of renewal and expansion of Cdk2-null bone marrow cells in a wild-type environment. We harvested Cdk2–/– and wild-type bone marrow and transplanted it into C57BL/6 lethally irradiated mice. The competitive repopulation was performed with similar numbers of host cells (Ly5.1) and donor Cdk2–/– or Cdk2+/+ cells (Ly5.2) to measure HSC function. Host and donor cell repopulation was determined by expression of the markers Ly5.1 and Ly5.2. We analyzed subpopulations of cells from bone marrow, thymus, spleen, and peripheral blood after more than 6 months to determine whether stem cell self-renewal and differentiation is maintained in the absence of Cdk2. We measured the Ly5.2/Ly5.1 ratios, which indicate the capacity of donor cells to establish multilineage cell reconstitution in competition with recipient cells. The repopulation capacity of Cdk2–/– transplanted cells was similar to that of Cdk2+/+ cells in bone marrow, thymus, spleen, and peripheral blood (Fig. 5C). This result suggests that the loss of Cdk2 in vivo does not affect the renewal of stem cells and the expansion of primitive progenitors, since they contribute to the same extent as wild-type cells to the reconstitution of hematopoietic subpopulations.

Cdk2 is a major kinase controlling S phase; however, our results indicated that Cdk2 functions are most likely compensated for by other Cdks. Cell cycle regulation required for maintenance of stem cells and expansion of primitive progenitors can occur in the absence of Cdk2.

Analysis of Cdk2-null splenocytes. The spleen is the lymphoid organ where clonal expansion of B and T cells that react to specific antigens takes place. This antigenic activation is mediated by antigen-presenting cells, such as macrophages, and induces the expression of multiple cytokines and reentry into cell cycle. From our thymus, bone marrow, and transplantation studies, we expected that the numbers of splenocytes would not be affected. Indeed, we observed similar numbers of cells in Cdk2+/+ ([74.5 ± 21] x 106 cells) and Cdk2–/– ([74.5 ± 18] x 106 cells) spleen. We analyzed different subpopulations (B cells, T cells, and macrophages) by flow cytometry and observed equivalent ratios of B lymphocytes (B220+), T lymphocytes (H57+/CD4+ or CD8+), and macrophages (Mac-1+/Gr-1+) in spleens from Cdk2–/– mice compared to their wild-type littermates (Fig. 6A). Among the T lymphocytes, the expression profiles of CD62L (positive in naive T cells) and CD44 (positive in memory T cells) were comparable in each CD4+ or CD8+ subpopulation (data not shown). As mentioned in the introduction, a good example of the central role of Cdk2 is its induction after TCR activation and the increased Cdk2 activity upon IL-2 stimulation (15, 29). This dual signal is mimicked in vitro by anti-CD3 and anti-CD28 antibodies, allowing quiescent T lymphocytes to enter into cell cycle. To determine whether Cdk2 affects T-lymphocyte activation, we measured the up-regulation of CD25, the IL-2 receptor (IL-2R), which is expressed on activated T cells and correlates with their capacity to proliferate. After 3 days of anti-CD3/CD28 stimulation, the expression levels of CD25 were increased compared to day 0 to the same extent in wild-type and Cdk2-null T lymphocytes, regardless of their CD4 or CD8 status (Fig. 6B). Moreover, the total numbers of CD4+ or CD8+ cells increased after stimulation in the same range, independently of genotype (Fig. 6C). The ratio of CD8+/CD4+ after anti-CD3/CD28 stimulation indicated that the CD8+ population expanded faster than the CD4+ population, leading to an inverted ratio at day 3 compared to unstimulated cells on day 0. However, we have not observed significant differences in these ratios between Cdk2–/– and Cdk2+/+ T lymphocytes (Fig. 6C). These results suggest that Cdk2 is not required in the signaling pathways induced by combined activation of TCR and IL-2R, which induces T lymphocytes to proliferate, enter S phase, and expand clonally. To confirm this result, we performed an in vivo antigenic stimulation experiment using ovalbumin (OVA) as the antigen. After two immunizations with OVA (2 weeks between each injection), spleens from Cdk2-null and wild-type mice were harvested and splenocytes were stimulated in culture in the presence of OVA. No significant differences in proliferation, analyzed by BrdU incorporation (an indicator of DNA replication), CD8+/CD4+ ratio, IL-2R expression, and gamma interferon expression were observed between Cdk2–/– and Cdk2+/+ T lymphocytes (data not shown).


Figure 6
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  		FIG. 6. Analysis of splenocytes in Cdk2–/– mice. (A) Analysis of splenocyte subpopulations by flow cytometry in Cdk2–/– mice (n = 4) compared to their wild-type littermates (n = 3). Dot plots and subpopulation percentages of representative data are shown. (B) Analysis of IL-2R (CD25) expression in CD4- or CD8-positive cells by flow cytometry. Lymphocytes harvested from Cdk2–/– (n = 12) or wild-type (n = 10) spleens were stimulated with anti-CD3 and anti-CD28 for 3 days. Histograms at day 0 and day 3 of a representative experiment are shown. (C) Expression of CD4 and CD8 receptors on unstimulated (day 0) or stimulated (day 3 after anti-CD3 and anti-CD28 treatment) lymphocytes was analyzed by flow cytometry. Total cell numbers of CD4+ or CD8+ lymphocytes were determined (percentage of population times spleen cellularity), and average values from three independent experiments are plotted. CD8/CD4 ratios are indicated on the graph. (D) CBA was performed with cell culture supernatants from LPS-stimulated macrophages to measure cytokine gene expression. Average values of cytokine expression in Cdk2–/– (n = 2) or Cdk2+/+ (n = 2) cells are plotted.

We have shown that macrophages are normally represented in bone marrow and spleen of Cdk2–/– mice. To determine whether macrophage stimulation was affected, we decided to harvested bone marrow cells and expand them in culture under macrophage differentiation conditions. After 6 days in culture in the presence of G-CSF, we stimulated adherent cells with different doses of LPS in the presence of gamma interferon to induce inflammatory cytokines such as tumor necrosis factor alpha (TNF-{alpha}), IL-10, IL-6, and monocyte chemoattractant protein 1 (MCP-1). After 24 h of stimulation, a CBA analysis (see Materials and Methods) of the supernatant of these macrophages was performed to analyze the cytokine expression profile. We also determined the expression of Mac-1- and F4/80-specific macrophage markers by flow cytometry and found similar macrophage differentiation rates in cultures from both genotypes (Cdk2+/+, 46.4% ± 7%; Cdk2–/–, 46.1% ± 6% of Mac-1+; F4/80+). The induction profiles of cytokines from Cdk2–/– and Cdk2+/+ macrophages were comparable for IL-10, and we observed slight decreases in TNF-{alpha}, IL-6, and MCP-1 expression, which are probably not statistically significant (Fig. 6D). The present results suggest that upon stimulation, the inflammatory response associated with macrophages is not impaired in Cdk2 knockout mice.

We conclude from our analysis of splenocytes that macrophages and T lymphocytes are properly activated upon antigen stimulation in the absence of Cdk2. Moreover, their expansion and differentiation, which involve entry or exit of the cell cycle, do not seem to be impaired.


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DISCUSSION
 
Cdk2 knockout mice are an excellent model for in vivo study of the consequences of complete Cdk2 inactivation. Although RNA interference and pharmacological studies are very useful, they do not achieve the high specificity of knockout mice. Numerous in vitro studies of hematopoietic and immune cells have suggested Cdk2 as a downstream target of signaling pathways associated with mitogenic cytokines or antigen activation. In this context, two roles for Cdk2 have been characterized: one is related to cell cycle progression and S phase entry; the second, less expected, is the requirement for thymocyte apoptosis. Both functions are essential for proper homeostasis of hematopoietic and immune cells, since it affects their self-renewal capacity, maturation, and ability to respond to antigens. Through biological assays, we addressed the question of the in vivo role of Cdk2. Surprisingly, we found that Cdk2 was not required for apoptosis associated with antigen-induced negative selection of thymocytes, as previously described by pharmacological in vitro studies. We also performed a broad range of assays to determine whether other cell types in bone marrow and spleen would be affected by the lack of Cdk2. It appears that Cdk2 functions are most likely compensated for, since we did not observe any significant differences between Cdk2–/– and wild-type animals.

Cdk2 activity has been suggested to contribute to apoptosis in different cell types. The Cdk inhibitor roscovitine has been determined to be able to block apoptosis in thymocytes after irradiation, dexamethasone, heat shock, etoposide, PMA, or anti-CD3 treatment but not anti-Fas/CD95 treatment (17, 20). Roscovitine protects thymocytes from apoptosis, and an apparent link between apoptosis and increase in Cdk2 activity has been observed. Moreover, roscovitine blocks the apoptotic pathway independent of TCR rearrangements (35), suggesting a role independent of DNA repair, where Cdk2 could also be involved (28). Our results indicate that roscovitine has the same effect in Cdk2-null thymocytes, which indicates that Cdk2 is not the main target of roscovitine under these conditions. Recently, another study has reported that roscovitine induces apoptosis in neutrophils (32). These nonproliferating cells express Cdk2, suggesting that this kinase may play a role unrelated to the cell cycle. However, we did not investigate how the lack of Cdk2 affects the survival of neutrophils and how roscovitine might affect neutrophils through other targets and pathways. It is possible that roscovitine targets other Cdks, compensating for Cdk2. Among the potential Cdks, we can rule out Cdk3, the closest homologue of Cdk2, since C57BL/6 mice carry a premature stop codon in the kinase domain (36). Hakem et al. (20) showed that Cdk4 and Cdk1 kinase activities were not induced by dexamethasone treatment. We confirmed by genetic means that Cdk4 is not involved in this pathway. Furthermore, a Cdk1 inhibitor did not have the same effect as roscovitine, indicating that Cdk1 inhibition cannot protect thymocytes from apoptosis. We have shown that Cdk1 compensates for Cdk2 in vivo (2), but our present results show that Cdk1 is most likely not the target of roscovitine. Indeed, pharmacological inhibition of Cdk1 in Cdk2–/– thymocytes did not increase the survival rate after dexamethasone treatment. It was further confirmed that roscovitine treatment did not affect Cdk1 and cyclin B activity (see Fig. 2C). Neither is Cdk6 a good candidate, since Cdk2 and Cdk6 double knockout mice do not exhibit any phenotype associated with thymic maturation (25). Cdk5 and Cdk9 could potentially play a role, as these Cdks have been involved in apoptotic pathways and are inhibited by roscovitine (9, 18). However, neither of these kinases has been associated with thymocytes so far, and further investigations are required to determine which proteins is the target(s) of roscovitine in thymocytes, preventing apoptosis.

Our results demonstrate that the in vivo functions of Cdk2 are compensated for in hematopoietic and immune cells. Most importantly, we found that although mouse embryonic fibroblasts display minor growth defects in short-term culture and Cdk2–/– mice are smaller than wild-type mice, long-term growth in a competitive graft experiment was not affected (see Fig. 5). Cell cycle regulation and control of proliferation are not dependent on Cdk2 (5, 31), which most likely explains the absence of phenotypes. Our study provides an extensive analysis of the hematopoietic system and confirms that Cdk2 is fully compensated for in vivo, even when cells are challenged by antigens or mitogenic signals. This indicates that knockout models are a complement to pharmacological studies in order to be able to determine the in vivo targets of a drug. The therapeutic value of a drug, in particular roscovitine, remains valid, but further detailed investigations are necessary to determine the complete biological effects. We should keep in mind that acute inhibition associated with drugs is different from embryonic inactivation. However, if embryonic plasticity allow the cells to compensate for Cdk2, it is likely that tumor cells may acquire a similar compensatory mechanism (6).


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ACKNOWLEDGMENTS
 
We thank Daniel McVicar, Nancy Jenkins, and Neal Copeland for advice, discussion, reagents, and support. We also thank Michael Sanford, Jeff Subleski, and Kim Klarmann for their contributions and Mary Beth Hilton, Matt McCollum, and Angie Smith for expert animal care. We are grateful to Daniel McVicar and Stephen Anderson for providing equipment and reagents. We really appreciated the help of the Kaldis laboratory. For comments on the manuscript, we thank Jonathan Ashwell.

This research was supported by the Intramural Research Program of the NIH, National Cancer Institute, Center for Cancer Research.


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FOOTNOTES
 
* Corresponding author. Mailing address: Mouse Cancer Genetics Program, Center for Cancer Research, National Cancer Institute—Frederick, Bldg. 560/22-56, 1050 Boyles Street, Frederick, MD 21702-1201. Phone: (301) 846-1988. Fax: (301) 846-7017. E-mail: kaldis{at}ncifcrf.gov Back

{triangledown} Published ahead of print on 7 May 2007. Back

{dagger} Present address: Oncodesign, Dijon 21076, France. Back

{ddagger} Present address: MedImmune Inc., Gaithersburg, MD 20878. Back


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Molecular and Cellular Biology, July 2007, p. 5079-5089, Vol. 27, No. 14
0270-7306/07/$08.00+0     doi:10.1128/MCB.00029-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.



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