Dissociation of CDK2 from Cyclin A in Response to the Topoisomerase II Inhibitor Etoposide in v-src-Transformed but Not Normal NIH 3T3 Cells

Experimental Cell Research 249, 327–336 (1999) Article ID excr.1999.4484, available online at http://www.idealibrary.com on

Dissociation of CDK2 from Cyclin A in Response to the Topoisomerase II Inhibitor Etoposide in v-src-Transformed but Not Normal NIH 3T3 Cells Guan Chen 1 and Masahiro Hitomi The Lerner Research Institute, Department of Molecular Biology, Cleveland Clinic Foundation, Cleveland, Ohio 44195

Our previous work has demonstrated that treatment of NIH 3T3 cells with etoposide (VP16), an inhibitor of DNA topoisomerase II and widely used anticancer agent, results in G2/M-phase arrest, whereas treatment of cells transformed by v-src, v-ras, or v-raf results in an S-phase blockage. The present studies describe the mechanistic aspects of this selective S-phase arrest in the v-src-transformed cells. The Sphase arrest in these cells was found to be coupled with depletion of cyclin A-dependent kinase activity. This decrease could not be explained by changes in the overall level of cyclin A, CDK2, p27, or p21 proteins. Rather, it was associated with a time-dependent reduction of CDK2 protein complexed with cyclin A following VP16 treatment. It was further shown that the decrease of cyclin A-associated CDK2 was linked to an increase of CDK2 protein in cyclin E immunocomplexes, which suggests that CDK2 might become redistributed following treatment with VP16. Thus, oncogenic transformation by v-src can trigger separation of CDK2 protein from cyclin A in response to VP16. This might contribute to the depletion of cyclin A-dependent kinase activity and the selective S-phase arrest by VP16 in v-src-transformed cells. © 1999 Academic Press Key Words: oncogenic transformation; S-phase arrest; etoposide; CDK2; cyclin A-dependent kinase.

INTRODUCTION

Progression through several major checkpoints in the cell cycle is controlled by multiple protein kinases, each of which contains a regulatory cyclin and a catalytic cyclin-dependent kinase (CDK) 2 [1–5]. The activity of these kinases is regulated by the expression level of each component, its phosphorylation status, and the 1 To whom correspondence and reprint requests should be addressed at Department of Molecular Biology, NC20, The Cleveland Clinic Foundation, 9500 Euclid Avenue, Cleveland, OH 44195. Fax: 216-444-0512. E-mail: [email protected]. 2 Abbreviations used: CDK, cyclin-dependent kinase; p21, Waf1 or Cip1; p27, Kip1; VP16, etoposide; topo IIa, DNA topoisomerase IIa.

presence of specific CDK inhibitory proteins [2]. In mammalian cells, the D cyclins with their catalytic partners CDK4 and CDK6 function as the cells leave G0 and progress through G1, while cyclin E/CDK2 is activated from G1 into S phase. Cyclin A/CDK2 operates in S and G2 phases, whereas cyclin B/CDK1 orchestrates the G2/M transition [2]. p21 (Cip1, Waf1), p27 (Kip1), and p57 (Kip2) belong to a category of broad-specificity inhibitors of cyclin/CDK complexes. But each of the Ink family members (p15, p16, p18, and p19) binds directly to CDK4 and CDK6 and functions as a specific inhibitor of the cyclin D-dependent kinases [1, 2]. It is becoming clear that many external signals including both those that stimulate growth, such as growth factors, and those that inhibit growth, such as DNA damaging agents, control cell proliferation through regulating the cell cycle. Thus, elucidating the machinery of cell cycle progression and its regulation by these signals is essential for understanding and controlling cell proliferation. While mitogenic growth factors and oncogenes have been found to trigger cell cycle progression by controlling progression through G1 phase [2, 6 –10], cellular response to DNA damage may occur at any phase of cell cycle [11–13]. In many cell types DNA damage response pathways cause arrest by regulating CDKcyclin via checkpoint proteins which sense damage and transduce an inhibitory signal [14, 15]. Checkpoints are regulatory pathways that control the order and timing of cell cycle transitions and which ensure the fidelity of genomic replication and separation at mitosis. Many checkpoints become activated by DNA damage. This results in an arrest in the cell cycle which provides time for DNA repair. In addition, this often leads to apoptosis [14, 15]. Recent studies demonstrated that one of the molecules participating in DNA damage-induced G2 arrest is CDK1 (CDC2), which is activated following dephosphorylation by Cdc25 [13, 16 –18]. The phosphatase Cdc25 appears to become active once it is phosphorylated on Ser 216 by protein kinase Chk1 following DNA damage [13, 16 –18]. These results have broadened our knowledge of the check-

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point signals interacting with the machinery that controls progression from the G2 to the M phase. It is possible, however, that different checkpoints could be activated following the same DNA-damaging treatment in normal and malignant cells, as the malignant cells can grow under conditions in which the normal ones are blocked [19]. If so, analyses of this difference, especially the responses to commonly used anticancer drugs, may be critical for understanding molecular mechanisms of the selectivity of chemotherapy in tumor cells and thereby may contribute to the development of novel therapeutic strategies by targeting tumor-specific checkpoints. Proliferating cells are vulnerable to DNA damage due to the demands of cellular growth and division [12]. As proliferative signaling is constitutively active in the tumor [20], it is reasonable to assume that the higher sensitivity of malignant cells to DNA-damaging agents is caused by these signals that are able to regulate checkpoint responses. Indeed, recent studies demonstrated that oncogenic transformation or overexpressing of a transcriptional factor E2F1 causes cells arrested in S phase in response to etoposide (VP16), a DNA topoisomerase II inhibitor and an chemotherapeutic agent widely used in the clinic [21–23]. In this report, mechanistic aspects of selective S-phase arrest induced by VP16 were investigated in v-src-transformed NIH 3T3 cells in comparison with nontransformed normal NIH 3T3 cells. Our results showed that this selective arrest is associated with loss of cyclin A-dependent kinase activity upon VP16 treatment. Further evidence demonstrates that this depletion can be mostly explained by a decrease of CDK2 protein in cyclin A immunoprecipitates and its increase in cyclin E complexes. These results, thus, revealed a novel mechanism for src oncogene-mediated activation of an S phase checkpoint, which may provide explanations for the selective toxicity in these and probably other malignant cells. MATERIALS AND METHODS Materials. Dulbecco’s modified Eagle’s medium (DMEM) and calf serum were obtained from Celox (Hopkins, MN) and Sigma (St. Louis, MO), respectively. All other materials for cell culture were supplied by Gibco (Grand Island, NY). Rabbit anti-cyclin A, anticyclin E, anti-p27, anti-CDK2, and anti-CDK4 polyclonal antibody; mouse anti-cyclin D1, anti-CDK1, anti-cyclin B1, and anti-actin; and goat anti-p21 monoclonal antibody were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). Anti-rabbit, anti-mouse, or antigoat IgG conjugated with horseradish peroxidase (HRP) from Boehringer Mannheim (Indianapolis, IN) was used as a second antibody at 1:2000 dilution. GST-Rb from Santa Cruz Biotechnology was used as substrate for CDK4 kinase determination. Renaissance Western blot chemiluminescence reagent and reflection autoradiography films from DuPont NEN (Boston, MA) were used for detection of signals. Protein A–Sepharose 4B was obtained from Zymed (San Francisco, CA). [g- 32P]ATP was provided by ICN (Costa Mesa, CA) at a specific activity of 4500 Ci/mmol. All other chemicals were from Sigma

Chemical Co. Etoposide solution (20 mg/ml) was purchased from Bristol-Myers Squibb Co. (Princeton, NJ). Cell lines and treatment with etoposide. NIH3T3 (3T3) cells and their subline stably transformed by viral oncogene v-src (Src) have been maintained in DMEM containing 10% calf serum and antibiotics at 37°C under the atmosphere of 5% CO 2. The sources and properties of these cells have been previously described [6, 23, 24]. Cells were seeded at 5 3 10 5 in a 100-mm petri dish with 10 ml growth medium and treated on the third day with 10 mM VP16 or vehicle control for various times as indicated. At the end of the treatment cells both in suspension and attached to the plate were washed twice with PBS and collected for different analysis. Western blotting and kinase assay. Cells were lysed for 20 min on ice in lysis buffer (50 mM Hepes, pH 7.6, 200 mM NaCl, 1 mM EDTA, 0.5% NP-40), which was supplemented with phosphatase inhibitors (5 mM b-glycerophosphate, 10 mM NaF, 0.1 mM Na 3VO 4) and protease inhibitors (1 mM PMSF, 1 mg/ml of aprotinin and leupeptin and 10 mg/ml of benzamide) immediately before use [25, 26]. For CDK4 kinase determination, cells were lysed in 50 mM Hepes, pH 7.6, 150 mM NaCl, 1 mM EDTA, 2.5 mM EGTA, 1 mM DTT, 0.1% Tween 20, and 10% glycerol containing 10 mg/ml leupeptin, 1 mM PMSF, 10 mg/ml aprotinin, 10 mM NaF, 5 mM b-glycerophosphate, and 100 mM orthovanadate [27]. After centrifugation (13,000g for 10 min at 4°C) the supernatant was collected and used immediately for Western blotting, immunoprecipitation or stored at 220°C. The kinase activity was usually determined within 2 months after the sample was frozen. Protein concentrations were determined by Bio-Rad/DC protein assay reagent (Bio-Rad, Hercules, CA) with BSA dissolved in the same lysis buffer as standard. For direct Western blot analysis, an aliquot of 50 mg protein was heated at 100°C for 5 min after mixing with an equal volume of 23 SDS/PAGE sample buffer and loaded into a 10 to 12.5% acrylamide gel based on the molecular weight of the interested molecule. After electrophoresis, the gel was transferred to a nitrocellulose membrane (MSI, Westboro, MA) in transfer buffer (30 mM Tris, pH 8.8, 120 mM glycine, 15% methanol, and 0.05% SDS) on a semidry transfer apparatus from Bio-Rad. The membrane was blocked with 5% nonfat milk in PBS containing 0.05% Tween prior to antibody treatments. The protein of interest was visualized with chemiluminescence. The blot was then stripped in a buffer (62.5 mM Tris–HCl, pH 6.8, 2% SDS, 200 mM 2-mercaptoethanol) at 50°C for 30 min. After extensive washing, the blot was used again for probing the next molecule, beginning from blocking. For immunoprecipitation, 200 mg of protein was incubated with 2 mg of respective antibody at 4°C overnight in a rotating wheel. The immune complex formed was then precipitated by adding 30 ml of 50% protein A–Sepharose beads or the same amount of those precoated with rabbit anti-mouse IgG when the mouse monoclonal antibody was used. The coating was achieved by adding 0.3 ml of the IgG solution (1 mg/ml) to 2 vol of 50% of protein A–Sepharose, followed by incubation for 1 h at 4°C with constant mixing. The mixture was then incubated for an additional 1 h at 4°C. The agarose beads with bound kinases were washed two times in lysis buffer and two times in Hepes kinase buffer (50 mM Hepes, pH 7.6, 10 mM MgCl2, 1 mM dithiothreitol). Supernatant above the Sepharose beads was carefully removed and combined with 30 ml of kinase buffer containing 1 mM NaF, 10 mM b-glycerolphosphate, 0.1 mM Na 3VO 4, 50 mM ATP, and 5 mCi of [g- 32P]ATP containing 0.5 mg of histone H1 or GST-Rb (for CDK4 kinase assay). The reaction was initiated by incubation at 30°C for 30 min with occasional agitation. The reaction mixture was then mixed with equal volumes of 23 SDS/PAGE sample buffer and heated in a boiling water bath for 5 min before loading onto SDS/PAGE in duplicate. One gel was dried and analyzed in a phosphorimager for kinase activity. Another gel was transferred to a nitrocellulose membrane for detection of the molecules coimmunoprecipitated as described above.

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FIG. 1. Selective induction of S-phase arrest by VP16 in v-src-transformed cells. Cells were treated with 10 mM VP16 for different times as indicated and collected for flow-cytometric analysis. Similar results were described before in this and other transformed cell lines [23].

RESULTS

VP16 Caused Multiple Alterations of Cell-CycleRegulating Molecules in 3T3 and v-srcTransformed Cells Treatment with 10 mM VP16 resulted in a transient S-phase block (6 h) and thereafter a persistent G2/M-

phase arrest in normal NIH 3T3 cells, while this treatment led predominantly to an S-phase arrest in v-srctransformed NIH 3T3 cells (Src) (Fig. 1). This effect, which appeared to be specific for topoisomerase II inhibitors, could also be observed in v-ras- or v-raf-transformed NIH 3T3 cells [23] and in E2F1-overexpressing myeloid 32D.3 cells [22].

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CDK2 and CDK4 protein in either cell line. Although these data provide a general profile of the molecular alterations in the two cell lines, additional functional analyses are needed to understand how these changes contribute to different types of cell cycle arrest in response to VP16. V-src Transformation Triggered Separation of CDK2 from Cyclin A and Thereby Led to Depletion of Cyclin A-Dependent Kinase Activity in Response to VP16 CDK2 kinase activity was measured to determine whether this enzyme contributes to S-phase arrest in Src cells. The basal level of this kinase was similar in the transformed and the nontransformed cells (Fig. 4, 0 h after the treatment). Of interest, CDK2 kinase activity was decreased similarly in the two cell lines, although S-phase arrest was seen only in Src cells, indicating that factors in addition to this kinase must be responsible for controlling S-phase progression under these conditions. The decrease of CDK2 activity is not due to p27 induction, as this inhibitory protein remained relatively constant following the treatment. An increase of p21 protein in this complex may be FIG. 2. Effects of VP16 treatment on expression of cell-cycleregulating molecules in NIH 3T3 cells. Cells were treated with 10 mM VP16 for the indicated time and lysates were collected thereafter for Western blot analysis. 50 mg of protein was loaded per lane in SDS/PAGE, probed with different antibodies and visualized with ECL. Some filters were stripped and reprobed with separate antibodies but the equal loading was confirmed in every case by probing with mouse monoclonal anti-actin antibody. Similar results were obtained in a separate experiment.

V-src-transformed NIH 3T3 cells were next used in comparison with the parental NIH 3T3 cells to explore the potential mechanisms involved in the S-phase arrest. Western blot analysis showed alterations of multiple cell-cycle-regulating molecules. The most significant differences between NIH 3T3 and Src cells are: (i) CDK1 protein is decreased in both cell lines in a timedependent manner, though more substantial in NIH 3T3 cells (Figs. 2 and 3). (ii) The decrease of cyclin D1 protein induced by VP16 in the normal cells is specific (Fig. 2), as this was not observed in Src-transformed cells (Fig. 3). (iii) p21 (Waf1) was induced in both cell lines, but the maximal induction occurred at 24 h in 3T3 cells and at 6 h in Src cells [23 and data not shown]. p27 levels, on the other hand, were not altered significantly (Figs. 2 and 3). (iv) There was a different pattern of cyclin A protein alterations with the two cell types; a faster migrating component was seen beginning at 12 h after VP16 treatment in 3T3 but not in Src cells (Figs. 2 and 3). This might have been due to phosphorylation and/or to protein degradation. On the other hand, there were no detectable changes in total

FIG. 3. Effects of VP16 treatment on expression of cell-cycleregulating molecules in v-src-transformed NIH 3T3 cells. Cells were treated and prepared for Western blot as described in the legend to Fig. 1. Similar results were obtained in a separate experiment.

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in the normal cells (Fig. 5). The activity of this kinase was further reduced with an increase in the exposure time and was almost undetectable at 72 h in the transformed cells. In 3T3 cells, on the other hand, at least 20% of the kinase activity remained over the entire period of the treatment. Since cyclin A-dependent kinase is essential during S-phase progression [2], and since there was an eightfold increase in its activity in untreated Src cells, the depletion of this kinase activity following VP16 treatment might be in large part responsible for the S-phase arrest in these cells. The decrease in cyclin A-dependent kinase activity in NIH 3T3 cells might be due to p21 Waf1 induction in this complex, as there were no substantial change in p27Kip1 or CDK2 protein in the complex. In Src cells, on the other hand, kinase reduction (95%) at 6 h may be a consequence of both p21 induction and CDK2

FIG. 4. Changes of CDK2 kinase activity and its associated molecules after VP16 in 3T3 and Src cells. Cells were treated with VP16 for the times indicated and lysates were prepared as in Fig. 1. 200 mg of protein was incubated overnight with 1 mg of rabbit polyclonal anti-CDK2 antibody at 4°C and the resulting immunoprecipitates were subjected to in vitro kinase assay using histone H1 as a substrate as described under Materials and Methods. The supernatant of the kinase reactions was loaded into two separate SDS/PAGE gels: one was for phosphorimage scanning for the kinase activity, and the other was transferred into a nitrocellulose membrane and used for detection of CDK2-associated molecules. Similar results were obtained in a separate experiment. For this and subsequent kinase assays, the immunoprecipitation and the kinase reaction for both cell lines was performed in the same experiment and the phosphorylated substrates were also separated in the same gel for comparison.

important, as it correlated well with the reduction of the kinase activity at all the time points analyzed in both lines. Additionally, the decrease of CDK2 protein in Src cells at 24 h must also be an important factor in the reduction in kinase activity. Since CDK2 can complex with either cyclin A or cyclin E, which function during S-phase progression and G1 to S transition, respectively [4], the change in CDK2 kinase activity may indicate an alteration of cyclin A and/or cyclin E-dependent kinases. The basal activity of cyclin E- and cyclin A-dependent kinases was increased by five- and eightfold, respectively, in Src cells compared to NIH3T3 cells without VP16 treatment (0 h), indicating the important role each may play in maintaining the transforming phenotype (Figs. 5 and 6). Within 48 h of VP16 treatment, cyclin E-associated kinase activity remained relatively constant in both cell lines (Fig. 6), thus ruling out the possibility of direct involvement of this kinase in the selective S-phase arrest. Six hours after VP16, however, 95% of the cyclin A-dependent kinase activity was lost in Src cells, whereas 50% of the activity remained

FIG. 5. Time-dependent depletion of cyclin A-dependent kinase activity and decrease of cyclin A-associated CDK2 in Src cells after VP16. Cell lysates were collected at different times following the treatment. 200 mg of protein was incubated overnight with 1 mg of rabbit polyclonal anti-cyclin A antibody at 4°C. Cyclin A-dependent kinase was assayed in vitro using histone H1 as a substrate. The associated molecules in cyclin A immunoprecipitates were detected in Western blot analyses as described in the legend to Fig. 3. The change of cyclin A-dependent kinase activity as well as cyclin A-associated CDK2 protein was expressed as percentage of untreated controls (0 h after VP16) in 3T3 and SRC cells, respectively. Similar results were obtained in a separate experiment.

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FIG. 6. Increase of cyclin E-associated CDK2 protein by VP16. 200 mg protein was incubated with 1 mg rabbit polyclonal anti-cyclin E at 4°C overnight and cyclin E-dependent kinase activity was assayed in vitro using histone H1 as a substrate. Cyclin E-associated molecules were detected in Western blot analyses using the same precipitates and the increase of CDK2 protein in the complex was expressed as fold increase over control in 3T3 and SRC cells, respectively. Similar results were obtained in a separate experiment.

protein depletion (70%) in the complex. However, reduction of cyclin A-dependent kinase activity at 24 h (96%) was most likely due to the tremendous depletion of CDK2 protein from cyclin A complex in the transformed cells, since induction of p21 and/or p27 was not observed at these time points (Fig. 5). Therefore, these experiments demonstrate that the selective S-phase arrest in Src cells in response to VP16 is associated with the depletion of cyclin A-dependent kinase activity, which may result from the selective depletion of CDK2 protein from the cyclin A/CDK2 complex. VP16 Caused an Increase of CDK2 Protein in Cyclin E Complex in Both Cell Lines As no significant loss of total CDK2 protein was observed in Western blot analysis using the whole cell lysates (Fig. 3), experiments were carried out to investigate the mechanisms of CDK2 protein reduction in complexes with A complex in Src cells. Analyses of immunoprecipitates with cyclin E antibody were performed. Cyclin E-dependent kinase was not substantially altered in either cell line in response to VP16 treatment, although it was consistently higher in Src cells (Fig. 6). Interestingly, CDK2 protein complexed with cyclin E in the transformed cells actually increased about 2.5- to 4.0-fold following VP16 (Fig. 6). The levels of the two inhibitory proteins p21 and p27 complexed with cyclin E were also significantly in-

creased following treatment, but only in Src cells (Fig. 6). Although CDK2 complexed with cyclin E protein was also increased by VP16 in NIH 3T3 cells, increases were not as great and were not associated with compensatory increases of p21 and p27 protein. These experiments demonstrated that VP16 treatment can increase CDK2 protein in cyclin E complex in both NIH 3T3 and Src cells, but this increase is only associated with increases in p21 and p27 in the transformed cells. Since no significant alterations in total CDK2 protein within Src cells were observed by direct Western blot analyses of whole cell lysates (Fig. 3), and since the increase of CDK2 associated with cyclin E correlated with its depletion in the cyclin A complex (Fig. 5), these results suggest the possibility that a redistribution of CDK2 protein occurred from association with cyclin A to association with cyclin E following VP16 treatment, although further study of this phenomenon is required. V-src Transformation Did Not Affect CDK4 and CDK1 Kinase Activity in Response to VP16 Changes in cyclin-dependent kinases required during G1 and G2/M phases were also analyzed in order to rule out their potential involvement in the phenomenon described above. CDK4 kinase activity decreased similarly in both cell lines following VP16 treatment, with a 1.5-fold higher basal activity in Src cells (0 h, Fig. 7A), as was observed for CDK2 kinase (Fig. 4). The decrease of CDK4 kinase activity was not due to alterations in CDK4 protein or p27 protein, as there was only a slight decrease in CDK4 protein in Src cells at 48 and 72 h, and p27 actually decreased in both cell lines following treatment. The decreased kinase activity can, however, be largely explained by a decrease in the amount of cyclin D1 protein in the complex (Fig. 7A). It is of interest to note that the similar pattern of kinase change in these two cell lines following VP16 treatment was maintained by altering the catalytical and inhibitory subunits of the complex; i.e., the decrease in cyclin D1 protein was compensated for by the decreased content of inhibitory p27, while the amount of CDK4 protein remained relatively constant. The situation is clearly different than that observed with cyclin A- or cyclin E-dependent kinase (Figs. 5 and 6). Other inhibitory proteins were undetectable in the CDK4 immunocomplex. Finally, no significant alterations in the activity of the CDK1 kinase in either cell line was observed following VP16 treatment, although there was a decrease in the fast-migrating cyclin B1 protein band in direct Western analysis at the later time points in Src cells (Fig. 7B, and data not shown). Thus, CDK4 and CDK1 kinase were clearly not involved in the selective S-phase arrest observed in Src cells, although some of their components changed substantially.

ETOPOSIDE TRIGGERS DISSOCIATION OF CDK2 FROM CYCLIN A IN v-src-TRANSFORMED CELLS

FIG. 7. Similar alterations of CDK4 and CDK1 kinase activity by VP16 in 3T3 and Src cells. 200 mg protein was incubated overnight with 1 mg of either rabbit polyclonal anti-CDK4 (A) or anti-CDK1 (B) antibody at 4°C. The kinase activity was assayed in vitro by using GST-Rb as a substrate for CDK4 and histone H1 as a substrate for CDK1. CDK4-associated molecules were detected by Western blot analyses in the same precipitates as described in the legend to Fig. 3. Similar results were obtained in a separate experiment.

DISCUSSION

Cancer arises through genetic alterations of a normal cell, resulting in alterations in the processes that control cell growth [2]. Perhaps the most basic difference between normal and tumor cells is the constitutively proliferative signaling in the malignant tissues, which may be derived from activation of an oncogene and/or mutation of a tumor suppressor gene. The proliferative signals not only lead to uncontrolled growth but also determine how the cells respond to DNA damage. This was demonstrated by the activation of an S-phase checkpoint in cells transformed by oncogenes or overexpressing E2F1 [22, 23, 28 –32]. The present studies have revealed that one mechanism operative in this S-phase checkpoint activation is the depletion of cyclin A-dependent kinase as a result of CDK2 sequestration from its complex. Since one biological consequence of S-phase arrest is apoptosis [22, 29 –33], as seen in NIH 3T3 cells transformed by src and other oncogenes [23], these results would have important implications for understanding the selective toxicity of cancer chemotherapy against tumor cells. The S-phase arrest in Src cells in response to VP16 occurs as a result of an activation of the S-phase check-

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point. This is very likely mediated by depletion of cyclin A-dependent kinase activity, leading to deregulated E2F-1 activity. The following several lines of evidence support this assumption. (i) Cyclin A in a complex with CDK2 is not only required for efficient DNA replication, but is also essential for maintaining orderly S-phase progression [34 –38]. (ii) The cyclin A/CDK2 kinase binds directly to the transcription factor E2F-1 and phosphorylates its heterodimeric partner DP-1, thereby inhibiting the E2F DNA-binding activity [37]. (iii) Disruption of either cyclin A kinase binding sites on E2F-1 or mutation of the phosphorylation sites on DP1 results in cells arrested in S phase and apoptosis [39, 40, 41]. (iv) NIH 3T3 cells expressing a mutant E2F-1 that lacks the cyclin A/CDK2binding domain exhibit about a twofold increase in S-phase duration relative to the cells expressing wildtype E2F-1, and these cells are much more sensitive to killing by S-phase-specific anticancer drugs including VP16 [28]. (v) Overexpression of E2F-1 alone can also similarly result in premature accumulation of cells in S phase and/or apoptosis [22, 29 –32]. Thus, the diminished cyclin A kinase activity in Src cells after VP16 would fail to down-regulate E2F activity in a timely manner (as in the case of E2F overexpression or the loss of its binding to this complex), leading to cells arrested in S phase and increased likelihood of apoptosis [22, 23, 29, 32]. The depletion of cyclin A-dependent kinase activity in Src cells can be mostly ascribed to the decrease of the catalytic subunit CDK2 protein in the complex in response to VP16, although p21 induction may contribute some at earlier time points. Since this decrease (Fig. 5) is correlated with its increase in the cyclin E complex (Fig. 6), which is only observed in Src cells without significant change of total level of CDK2 protein, this phenomenon may be due to its redistribution, i.e., a shift of CDK2 from complexing with cyclin A to cyclin E. In normal situations, the amount of CDK2 protein does not significantly fluctuate during the cell cycle, while its kinase activity changes significantly, with peaks during DNA synthesis and right before mitosis [42]. In both cases the majority of the kinase activity is attributable to association of CDK2 with cyclin A, although the cyclin E/CDK2 complex appears first during the transition from G1 to S phase. After its interaction with cyclin A during progression into S phase, most of cyclin E is degraded [42]. Thus, src transformation caused a reversed flow of CDK2 protein in response to VP16, from its association with cyclin A to complexing with cyclin E (Figs. 5 and 6). Although redistribution of the inhibitory protein p21 and p27 has been reported among different cyclin–CDK complexes in cells after estradiol [43], UV irradiation [44], and TGF-b [45], our data here provide an example of CDK2 protein redistribution in response to VP16. This may

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present a novel mechanism by which cyclin A kinase activity is regulated with cyclin E as a reservoir for CDK2 protein in response to VP16. The CDK2 protein shift probably occurs as a result of decreasing affinity to cyclin A and/or increasing affinity to cyclin E. The exact mechanisms involved, however, remain to be explored further. V-src transformation is sufficient to cause CDK2 protein sequestration from cyclin A complex when challenged with VP16. This might explain why VP16 treatment causes normal cells to respond predominantly with a G2/M block [22, 23, 46 – 48], while oncogenic transformed cells [23] or cells overexpressing E2F-1 [22] are primarily arrested in S phase. Thus, the proliferative signals conferred by v-src provide cells with greater proliferating potential in normal situations, whereas they also activate the S-phase checkpoint by redistributing the CDK2 pool upon VP16 exposure, prior to the G2/M checkpoint as observed in normal diploid cells [23]. Previous work has demonstrated that the lack of p21 following treatment with anticancer drugs can lead to an uncoupling of S phase and mitosis, which is followed by apoptosis, thereby preventing cells with damaged DNA from reinitiating S phase [48]. This has been described as a consequence of the checkpoint failure [48]. Thus disruption of a normal cell cycle checkpoint should have dramatic effects on the response to anticancer drugs in mammalian cells. As defects in cell cycle checkpoints are one major characteristic of malignant cells [14], regulation of checkpoints based on these differences would accordingly correspond to a selective chemotherapeutic strategy. Nuclear DNA topo IIa, the proposed cellular target for VP16, may play an important role in activation of the S-phase checkpoints and the selective toxicity in E2F-1-overexpressing cells or oncogenic-transformed cells [22, 23]. Topo IIa is an essential nuclear enzyme that maintains proper DNA topology [49 –51]. Its expression is highly regulated in the cell cycle, i.e., lower in G1 and early S and higher in G2/M phase. Many transformed or tumor cells expressed higher levels of topo IIa, whereas it becomes lower in differentiated tissues [49 –56]. Our previous work demonstrated an increased amount of topo IIa protein in NIH 3T3 cells transformed by oncogene src, ras, or raf, and Ras signaling can directly stimulate its transcription through MEK/ERK pathway (Chen et al., submitted for publication). Furthermore, Ras signaling was found to induce topo IIa in all phases throughout the cell cycle (Stacey et al., submitted for publication). It is very likely that the elevation of topo IIa in Src cells may be involved in the S-phase checkpoint activation. That is, elevation of topo IIa may not only constitute an important nuclear component of the constitutive proliferating signaling pathways in normal situations in Src cells, but may also facilitate the preactivation of the

S-phase checkpoint in cooperation with or via regulation of cyclin A/CDK2 in response to the topo II inhibitor VP16. This speculation is further supported by the observation that the selective sensitization in E2F1overexpressing cells occurs only to the topo II inhibitor VP16 and adriamycin, but not to other anticancer drugs such as Taxol and 5-fluorouracil [22], and that E2F1 can also stimulate topo IIa promoter activity (Chen et al., unpublished). The detailed mechanisms for the interrelationship among oncogenes, topo IIa, and cell-cycle-regulating molecules warrants further investigation. We thank Dennis Stacey for helpful suggestions throughout this project and Nancy Wang for help in preparation of the manuscript. This work was supported in part by Grant GM 52271 to Dennis Stacey from the National Institutes of Health.

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