Antiprogestin Inhibition of Cell Cycle Progression in T-47D Breast Cancer Cells Is Accompanied by Induction of the Cyclin-Dependent Kinase Inhibitor p21

Elizabeth A. Musgrove, Christine S. L. Lee, Ann L. Cornish, Alex Swarbrick and Robert L. Sutherland

Cancer Research Program Garvan Institute of Medical Research St. Vincent’s Hospital Sydney, New South Wales 2010, Australia


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Progestin antagonists inhibit the proliferation of progesterone receptor-positive cells, including breast cancer cells, by G1 phase-specific actions, but the molecular targets involved are not defined. Reduced phosphorylation of pRB, a substrate for G1 cyclin-dependent kinases (CDKs) in vivo, was apparent after 9 h treatment of T-47D breast cancer cells with the antiprogestins RU 486 or ORG 31710, accompanying changes in S phase fraction. Although the abundance of cyclin D1, Cdk4, and Cdk6 did not decrease, cyclin D1-associated kinase activity was reduced by ~50% at 9–18 h. Similarly, cyclin E-associated kinase activity decreased by ~60% at 12–24 h in the absence of significant changes in the abundance of cyclin E and Cdk2. The CDK inhibitor p21 increased in mRNA and protein abundance and was present at increased levels in cyclin D1 and cyclin E complexes at times when their kinase activity was decreased. Increased p21 protein abundance was observed in another antiprogestin-sensitive cell line, BT 474, but not in two breast cancer cell lines insensitive to antiprogestins. These data suggest increased p21 abundance and concurrent inhibition of CDK activity as a mechanism for antiprogestin induction of growth arrest. Antiprogestin effects on proliferation were markedly reduced after ectopic expression of cyclin D1, indicating that inhibition of cyclin D1 function is a critical element in antiprogestin inhibition of proliferation. However, these data also implicate regulation of cyclin E function in antiprogestin regulation of cell cycle progression.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Cancers of steroid hormone target tissues, i.e. breast, prostate, endometrium, and ovary, account for a third of newly diagnosed cancers (1). Many of these cancers retain steroid responsiveness, leading to the development of synthetic steroid antagonists as potential therapeutic agents for hormone-dependent cancers. The success of this strategy is illustrated by the nonsteroidal antiestrogen tamoxifen, which is among the most effective specific therapies for breast cancer (2). In animal models of mammary cancer the antiprogestin RU 486 is as effective as tamoxifen in inhibition of tumor growth (3). Preliminary clinical trials of RU 486 in patients with metastatic breast cancer demonstrated some efficacy (4, 5), but frequent side effects, attributed to the potent antiglucocorticoid activity of RU 486, were observed (5). This problem may be minimized by more recently developed antiprogestins that display little antiglucocorticoid activity (6). Thus, antiprogestins may be of use in the treatment of breast cancer, in addition to their obstetric and gynecological uses (7), and their optimal clinical use is likely to be aided by a more detailed understanding of their molecular modes of action as anticancer agents.

Studies using breast cancer cells in vitro and rodent mammary tumors in vivo suggest that the antitumor activity of antiprogestins is mediated by inhibition of proliferation (8). Like other steroid or retinoid receptor ligands, antiprogestins have cell cycle phase-specific effects on cell proliferation (9, 10). Antiprogestin treatment leads to accumulation of breast cancer cells in G1 phase at the expense of cells in S, G2, and M phases (11, 12, 13), but the molecular targets involved in mediating this effect have not been defined. However, recent studies implicate regulation of cyclin/cyclin-dependent kinase (CDK) activity, particularly cyclin D-associated kinase activity, in steroidal regulation of proliferation. Cyclin D1 appears to be necessary for the progesterone-dependent development and differentiation of the mammary gland, because cyclin D1-deficient mice fail to develop lobular alveoli during pregnancy (14, 15), and this phenotype is shared by progesterone receptor-deficient mice (16). Furthermore, antiestrogen and retinoid inhibition of breast cancer cell cycle progression is accompanied by decreased cyclin D1 function (17, 18, 19), whereas decreased cyclin D3 function contributes to glucocorticoid inhibition of lymphoma cell proliferation (20).

Sequential activation of cyclin/CDK enzyme complexes regulates progress through the cell cycle. These enzymes are regulated at multiple levels, providing a variety of possible means by which overall activity of the complex, and hence the rate of cell cycle progression, might be modulated. The catalytic activity of CDKs depends not only on cyclin association but also on appropriate phosphorylation of the CDK subunit (21). The CDK-activating kinase (CAK) is itself a cyclin/CDK complex (cyclin H/Cdk7) regulated by phosphorylation. However, regulation of cyclin abundance governs much of the regulation of CDK activity during cell cycle progression and is a frequent response to treatment either by mitogens or inhibitors of cell proliferation (22, 23). Alteration of cyclin abundance is sufficient to alter the rate of cell cycle progression because overexpression of cyclins D or E accelerates cells through G1 and, conversely, inhibition of their function by antibody microinjection prevents entry into S phase (23).

A further means of regulating cyclin/CDK function is provided by endogenous low molecular weight proteins that physically associate with the cyclins, CDKs, or their complexes and inhibit CDK activity (24). A growing family of such inhibitors, for which p16INK4 is the prototype, selectively targets Cdk4 and Cdk6 (24). A second family of inhibitors, including p21 (WAF1, Cip1, Sdi1) and p27 (Kip1), are active against a wider range of cyclin/CDK complexes (24). The balance between levels of inhibitor and cyclin/CDK complexes is thought to set a threshold for activation of the kinase. The mechanism for inhibition of CDK activity has not been defined, but p21 immunoprecipitates display kinase activity indicating that association per se is not sufficient for inactivation (25, 26). Increased p21 or p27 abundance accompanies inhibition of proliferation during quiescence, senescence, and differentiation and contributes to inhibition of growth by transforming growth factor-ß (24, 27).

The retinoblastoma tumor suppressor protein, pRB, is a critical substrate for the G1 CDKs. pRB is hypophosphorylated during early G1 and in this form is growth-inhibitory. The pRB hyperphosphorylation that relieves this growth inhibition is first apparent in late G1 phase and continues during the remainder of the cell cycle (28). Cells without functional pRB lose dependence on cyclin D1 for G1 progression but demonstrate an absolute requirement for cyclin D1 upon reintroduction of pRB (29, 30). These data provide compelling evidence that pRB is a critical physiological target for cyclin D1. However, it is likely that cyclin E/Cdk2 also phosphorylates pRB in vivo, perhaps contributing to the further phosphorylation of pRB as cells progress into S phase (28).

In breast cancer cells cyclin D1 is both necessary and sufficient for G1 phase progression (31, 32). Furthermore, cyclin D1 abundance is rate-limiting in these cells as well as in fibroblasts (32, 33, 34). Initial studies demonstrated that neither cyclin D1 nor Cdk4 mRNA decreased in abundance after antiprogestin treatment, despite inhibition of proliferation (12, 19). However, as outlined above, other mechanisms for regulation of cyclin function exist and, in view of increasing evidence for regulation of cyclin function after treatment with steroids, steroid antagonists, and retinoids, the possibility that antiprogestins might regulate cyclin function in T-47D human breast cancer cells was investigated. Both RU 486, which is the prototypic progestin antagonist but also has glucocorticoid antagonist activity, and ORG 31710, which is representative of newer progestin antagonists with little antiglucocorticoid activity, were used. These studies demonstrate that antiprogestins induce p21 and regulate G1 cyclin function and indicate that this is likely to account for their inhibition of breast cancer cell proliferation.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
T-47D human breast cancer cells were cultured under conditions leading to optimal growth rates, i.e. in medium supplemented with insulin and FCS, to maximize sensitivity to growth inhibition. Initial experiments sought to define the effects of progestin antagonists on cell cycle progression under these conditions. Both RU 486 and ORG 31710 led to a concentration-dependent decrease in relative cell number (Fig. 1AGo). ORG 31710 was more potent than RU 486 such that the response to ORG 31710 was maximal at 1 nM, compared with 10 nM after RU 486 treatment (Fig. 1AGo). The decrease in cell number was associated with a sustained decrease in the proportion of cells in S phase over the same concentration range, apparent within 1 day of treatment and at concentrations of >=1 nM, maintained until the conclusion of the experiment at 5 days (Fig. 1BGo). Maximally effective concentrations of RU 486 (100 nM) or ORG 31710 (10 nM) were chosen on the basis of these data and used for further experiments.



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Figure 1. Effect of Antiprogestins on T-47D Cell Proliferation

A, Exponentially proliferating T-47D human breast cancer cells were treated with RU 486 ({circ}) or ORG 31710 (•) at the indicated concentrations. Proliferation was assayed after 6–7 days’ treatment using a colorimetric assay, and absorbance of the treated wells is presented relative to that in vehicle-treated control wells, as relative cell number. Data points indicate mean ± SEM of values from two or three replicate experiments each performed in quadruplicate or (points without error bars) mean of quadruplicates in a single experiment. B, Exponentially proliferating T-47D human breast cancer cells were treated with ethanol vehicle or the indicated concentrations of RU 486 or ORG 31710. Individual flasks were harvested and stained for DNA analysis by flow cytometry after 1 ({square} ), 3 (), or 5 days ({blacksquare}) treatment. Data represent mean ± SEM or range of results from two to four experiments. The S phase of control cells was similar over the entire experiment and, therefore, data from days 1, 3, and 5 have been pooled and are shown as mean ± SEM (C,). C, Exponentially proliferating T-47D human breast cancer cells were treated with ethanol vehicle (x) RU 486 (100 nM, {circ}), or ORG 31710 (10 nM, •). Individual flasks were then harvested and stained for DNA analysis by flow cytometry. Data represent mean ± SEM, where this is greater than the size of the symbol used, of at least three points from a total of seven experiments (RU 486) or five experiments (ORG 31710). The S phase fraction is significantly reduced compared with time-matched controls after 9–24 h treatment: P < 0.04 at 9 h for both compounds, P < 0.0001 for 12–24 h RU 486 treatment, and P < 0.001 for 12–24 h ORG 31710.

 
To characterize the initial decrease in S phase in more detail, cell cycle phase distribution was determined at intervals after antiprogestin treatment. The proportion of cells in S phase remained near control levels for 6 h but began to decline at 9 h and reached a minimum after 12–18 h (Fig. 1CGo). Although the time courses for the two compounds were similar, ORG 31710 was slightly more effective. The effects of both antiprogestins on S phase fraction decreased between 18–24 h exposure but were thereafter maintained (Fig. 1Go, B and C).

Since pRB is a physiological substrate for the G1 phase CDKs, pRB phosphorylation in vivo was examined to determine whether changes in CDK activity might accompany antiprogestin treatment. In untreated exponentially proliferating cells pRB was predominantly in the hyperphosphorylated form (ppRB, Fig. 2Go). After 9 h or more of antiprogestin treatment the abundance of hypophosphorylated pRB increased, reaching a more than 2-fold increase after 18 h ORG 31710 treatment, and there was a concomitant decrease in the abundance of ppRB (Fig. 2Go and data not shown). On average, the ppRB/pRB ratio was reduced to ~50% of control at 12–24 h (Fig. 2BGo).



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Figure 2. Effect of Antiprogestins on pRB Phosphorylation

Exponentially proliferating T-47D human breast cancer cells were treated with RU 486 (100 nM) or ORG 31710 (10 nM), and whole cell lysates were prepared at intervals. Equal amounts of protein were separated by SDS-PAGE, transferred to nitrocellulose, and blotted using a monoclonal antibody specific for pRB. A, Representative Western blot. Hyperphosphorylated pRB (ppRB) exhibits reduced electrophoretic mobility compared with hypophosphorylated pRB (pRB). B, The ratio between the optical densities of the ppRB and pRB bands is presented relative to the average of vehicle-treated controls. Data represent mean ± SEM from a total of six experiments. Control (X); RU 486 ({circ}); ORG 31710 (•). Ratio is significantly less than control; P < 0.02 for 9–24 h RU 486 treatment; P = 0.05 for 9 h ORG 31710 treatment; and P < 0.01 for 12 or 18 h ORG 31710 treatment.

 
To identify changes in CDK activity that might be responsible for the decreased pRB phosphorylation, immunoprecipitates from cells treated with antiprogestin were used in kinase assays in vitro. Cyclin D1-associated kinase activity was measured using a pRB fusion protein substrate. The specificity of this assay has been demonstrated as follows: kinase activity was inhibited by addition of either recombinant GST-p16INK4 or GST-p21, was directed toward pRB fusion proteins but not histone H1, and was increased after ectopic cyclin D1 expression (Ref. 35 and B. Sarcevic, unpublished data). Kinase activity decreased significantly to reach a minimum of ~50% after 9–18 h ORG 31710 treatment (Fig. 3AGo). This decrease, while modest, was statistically significant at 9 and 12 h. Cyclin E-associated kinase activity was measured using histone H1 substrate and was decreased by ~60% from 12–24 h ORG 31710 treatment (Fig. 3BGo). RU 486 treatment led to similar effects on kinase activity (not shown). These data demonstrated a decrease in G1 cyclin-associated kinase activity which paralleled decreased pRB phosphorylation.



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Figure 3. ORG 31710 Effects on Cyclin D1-Associated and Cyclin E-Associated Kinase Activity

Exponentially proliferating T-47D human breast cancer cells were treated with ORG 31710 (10 nM), and cell lysates were prepared at intervals. A, Kinase activity of cyclin D1 immunoprecipitates was measured using a pRB fusion protein substrate. A representative autoradiogram is shown. Graph shows mean ± SEM of data pooled from three experiments after background subtraction as described in the Materials and Methods. Hatched bar indicates SEM of pooled control data. Activity is significantly less than control at 9 h (P = 0.0075) and 12 h (P = 0.046). B, Kinase activity of cyclin E immunoprecipitates was measured using histone H1 substrate. A representative autoradiogram is shown. Graph shows mean of data pooled from duplicate experiments. Range is shown where greater than the size of the symbol used. Hatched bar indicates SEM of pooled control data. Activity is significantly less than control from 6–24 h (P < 0.05 at 12 and 18 h; P < 0.0025 at 6 and 24 h).

 
Cyclin and CDK abundance were examined by Western blotting to identify possible mechanisms underlying the changes in CDK activity. Cyclin D1 protein abundance did not decrease after antiprogestin treatment but rather was either unchanged or slightly increased in abundance (Fig. 4AGo), consistent with previous examination of cyclin D1 mRNA after RU486 treatment (12, 19). No change in the abundance of cyclin E, Cdk4, or Cdk6 was detected (Fig. 4Go), and there was at most a minor decrease in Cdk2 abundance (not shown). Since regulation of the abundance of these cyclins and CDKs did not appear to contribute to decreased kinase activity CDK inhibitor expression was next examined. No change in the abundance of p27 was detected, but p21 levels increased markedly after 12 h or more antiprogestin treatment (Fig. 4BGo). Quantification of data from multiple experiments showed an average 4- to 5-fold increase after ORG 31710 treatment and a slightly smaller increase of >3-fold after RU 486 treatment (Fig. 4CGo). To determine the basis for the increase in p21 abundance Northern analysis was performed after ORG 31710 treatment. Within 6 h treatment p21 mRNA increased by ~3-fold and the increase was maintained at 18 h (Fig. 5Go), indicating that antiprogestin regulation of p21 mRNA levels is one cause of the increase in p21 protein levels.



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Figure 4. Antiprogestin Effects on Cyclin, CDK, and CDK Inhibitor Abundance

Exponentially proliferating T-47D human breast cancer cells were treated with RU 486 (100 nM) or ORG 31710 (10 nM), and cell lysates were prepared at intervals. Equal amounts of protein were separated by SDS-PAGE and transferred to nitrocellulose (A, B). Data in panel B were obtained by sequential incubation of the same blots with the indicated primary antibodies. C, Graph presents densitometric analysis of Western blots as the mean ± SEM of three to five experiments. RU 486 ({circ}); ORG 31710 (•). The abundance of p21 is significantly increased from 6–24 h RU 486 treatment (P < 0.05) and from 12–24 h ORG 31710 treatment (P < 0.02).

 


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Figure 5. ORG 31710 Effects on p21 mRNA Expression

Exponentially proliferating T-47D human breast cancer cells were treated with ORG 31710 (10 nM) for the indicated times and total RNA was extracted for Northern analysis.

 
To ascertain whether the abundance of p21 in cyclin immunoprecipitates increased as its abundance increased, cyclin D1 and cyclin E immunoprecipitates were examined. Cyclin D1 immunoprecipitates from cells treated with antiprogestin for up to 24 h contained cyclin D1, Cdk4, and p27 in amounts similar to those in control cells (data not shown). The amount of p21 coimmunoprecipitating with cyclin D1 was increased after >=12 treatment (Fig. 6AGo), to an average of ~3-fold relative to control. Cyclin E immunoprecipitates contained similar amounts of cyclin E and Cdk2 in control and antiprogestin-treated cells, but the amount of coimmunoprecipitated p21 increased by 2- to 3.3-fold after 12–24 h ORG 31710 treatment (Fig. 6BGo), when the kinase activity was decreased (Fig. 3BGo). Overall, these data are consistent with the interpretation that increased p21 abundance contributes to the reduction in cyclin-associated kinase activity after antiprogestin treatment.



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Figure 6. ORG 31710 Effects on Cyclin D1 and Cyclin E Complex Composition

Exponentially proliferating T-47D human breast cancer cells were treated with ORG 31710 (10 nM) and cell lysates were prepared. A, Cyclin D1 immunoprecipitates immunoblotted for p21. B, Cyclin E immunoprecipitates were immunoblotted sequentially for p21 and Cdk2.

 
To determine whether antiprogestin induction of p21 was always associated with inhibition of proliferation the effects of antiprogestin treatment were examined in several breast cancer cell lines expressing a range of progesterone receptor levels. MDA-MB-231 cells are progesterone receptor negative, while MCF-7 cells express 9.1 x 104 sites per cell and BT 474 cells express 4.5 x 105 sites per cell, in comparison with T-47D, which express 2.3 x 106 sites per cell (36). In MCF-7 or MDA-MB-231 cells no significant effect of antiprogestin was observed on the S phase fraction after 18 h treatment, when maximal effects were observed in T-47D cells (Fig. 7AGo). Similarly, there was no effect on cyclin D1, p21, or p27 abundance, nor was there any alteration in the amount of p21 coimmunoprecipitating with cyclin D1 (Fig. 7Go, B and C). In BT 474 no effect on S phase was observed after 15 h treatment but a significant, albeit small, effect was observed after ~30 h (Fig. 7AGo). This delay in action is likely a reflection of the longer doubling time of these cells, ~3 days compared with 1–1.5 days for the other three cell lines. Concomitant with the decrease in S phase fraction, there was an approximately 3-fold increase in p21 abundance but no change in the abundance of cyclin D1 or p27 (Fig. 7BGo). Cyclin D1 immunoprecipitates indicated increased p21 bound to cyclin D1 after ORG 31710 treatment of BT 474 (Fig. 7CGo). However, the level of p21 in cyclin D1 immunoprecipitates from treated BT 474 cells was still markedly lower than that in immunoprecipitates of a similar amount of cyclin D1 from T-47D cells. These data indicate an association between antiprogestin induction of p21 and inhibition of cell proliferation.



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Figure 7. ORG 31710 Effects on MCF-7, MDA-MB-231, and BT 474 Cells

Exponentially proliferating MCF-7, MDA-MB-231, and BT 474 human breast cancer cells were treated with vehicle or ORG 31710 (100 nM) for 18 h (MCF-7, MDA-MB-231) or 30 h (BT 474) unless otherwise indicated. A, S phase fraction was determined by flow cytometry. BT 474 data represent mean ± SEM of flasks harvested after 24–39 h treatment in two independent experiments. B, Cell lysates were prepared for immunoblotting. For each cell line, data were obtained by sequential incubation of a single filter with the indicated primary antibodies. C, Cyclin D1 immunoprecipitates immunoblotted for cyclin D1, p21, and Cdk4. Exponentially growing T-47D cells were immunoprecipitated and blotted in parallel for comparison. The amount of cellular protein immunoprecipitated was adjusted for each cell line to yield comparable amounts of immunoprecipitated cyclin D1. Data were obtained by sequential incubation of one filter with the indicated primary antibodies.

 
The data presented above suggest that decreased CDK activity after antiprogestin treatment might result from induction of p21 and subsequent association with cyclin/CDK complexes. However, the effect of the observed increase in p21 on kinase activity is likely to depend on the initial p21 occupancy of the cyclin/CDK complexes. Immunoblotting of cyclin D1 immunoprecipitates in parallel with cyclin D1-depleted lysate and mock-depleted lysate revealed that approximately a third of the total cellular p21 was associated with cyclin D1 in T-47D cells (Fig. 8AGo), suggesting that the cyclin D1 complexes in control cells contained significant amounts of p21. To assess the degree of saturation of the complexes, the effect of adding recombinant GST-p21 fusion protein to a constant amount of cyclin D1 immunoprecipitate from exponentially proliferating cells was determined. Recruitment of GST-p21 into the cyclin D1 complexes increased in proportion to the amount of GST-p21 added until the total p21 (the sum of the endogenous p21 and GST-p21) reached approximately 3 times the initial abundance (Fig. 8Go, B and C). However, increasing the amount of added GST-p21 by 10-fold led to a relatively modest further increase in the total p21 bound (Fig. 8Go, B and C). Thus the cyclin D1 complexes became saturated with p21 upon a 4-fold increase in associated p21. These data indicate that the ~3-fold relative increase in cyclin D1-associated p21 after antiprogestin treatment is likely to be sufficient to account for the observed decrease in kinase activity.



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Figure 8. Association of p21 with Cyclin D1 Complexes

A, Precleared lysates of exponentially proliferating T-47D cells were immunoprecipitated using either cyclin D1 antiserum linked to protein A beads (D1) or protein A beads alone (Con). Aliquots of the supernatants (Sup.) or immunoprecipitated protein (Pellet) were immunoblotted. Each lane contains protein derived from an equal amount of cellular protein. B and C, Increasing amounts of recombinant GST-p21 fusion protein were added to cyclin D1 immunoprecipitates of 750 µg lysate from exponentially proliferating T-47D cells. After 1 h at 30 C to allow association, the resulting complexes were immunoblotted (B). The intensities of the endogenous and exogenous p21 bands were added to yield total cyclin D1-associated p21, which is presented relative to the amount of endogenous p21 (C).

 
Since the activity of both cyclin D1- and cyclin E-associated CDKs was decreased, further experiments were designed to test the relative contributions of these responses to growth inhibition. T-47D cells transfected with a metal-responsive cyclin D1 construct, T-47D {Delta}MTcycD1-3 (32, 35), were used to examine the effects of increased cyclin D1 abundance on the response to antiprogestins. After zinc treatment of these cells cyclin D1 protein abundance increases within 3 h and reaches maximum levels by 6–9 h (32). The resulting acceleration of cells through G1 phase leads to an increase in the proportion of cells in S phase, which is maximal at 15–24 h (32). Since zinc induction of cyclin D1 and antiprogestin induction of p21 occur over a similar time course, the cyclin D1/p21 ratio would be expected to remain relatively constant after simultaneous treatment with ZnSO4 and antiprogestin. The increase in cyclin D1 abundance after zinc treatment was unaffected by simultaneous treatment with ORG 31710 (Fig. 9AGo). ORG 31710 treatment reduced the S phase fraction relative to untreated control cells when cyclin D1 levels were increased by up to ~2-fold (i.e. zinc concentrations of 30 µM or below) (Fig. 9BGo). A 2.5-fold increase in cyclin D1 abundance after treatment with 40 µM zinc prevented ORG 31710 treatment from decreasing the S phase fraction relative to untreated control cells, while a further increase in cyclin D1 abundance led to increased S phase despite the presence of antiprogestin (Fig. 9BGo). In the same experimental design, treatment of parental T-47D cells with up to 50 µM zinc did not significantly increase either cyclin D1 abundance or S phase fraction.



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Figure 9. Effect of Zinc Induction of Cyclin D1 in Antiprogestin-Treated Cells

T-47D {Delta}MTcycD1-3 cells proliferating exponentially were treated with the indicated concentration of ZnSO4 in the presence of either 10 nM ORG 31710 or 0.1% ethanol (control). A, Cells were harvested after 15 h treatment, and cyclin D1 abundance was determined by immunoblotting. Control ({circ}); ORG 31710 (•). B, Cells were harvested after 15 h treatment with ZnSO4 and ORG 31710 (•) and S phase fraction determined by flow cytometry. Points represent individual determinations in one of three experiments with similar results. The S phase of control cultures treated with 0.1% ethanol is indicated ({circ}– – –{circ}). C, Relationship between S phase fraction and cyclin D1 abundance in cells treated with 0–50 µM ZnSO4 in the presence of either 0.1% ethanol ({circ}) or ORG 31710 (•).

 
The S phase fraction of T-47D {Delta}MTcycD1-3 cells treated with zinc together with antiprogestin was consistently lower than that of cells treated with zinc in the absence of antiprogestin treatment, despite equivalent cyclin D1 levels (Fig. 9Go, A and B). However, consistent with previous data (32), cyclin D1 abundance and S phase fraction were linearly related in both cases (Fig. 9CGo). Lines of best fit generated by linear regression were essentially parallel, with slopes of 1.52 and 1.59 for control and ORG 31710, respectively (Fig. 9CGo), i.e. in either the presence or absence of antiprogestin a given absolute increase in cyclin D1 abundance led to the same absolute increase in S phase. Thus, the effect of antiprogestin treatment was to increase the amount of cyclin D1 necessary to achieve a particular S phase value, as would be expected if the threshold for cyclin D1 function had been increased as a result of p21 induction after antiprogestin treatment.


    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
The cell cycle-phase specific effects of steroids and steroid antagonists (10) suggest that their molecular targets include genes involved in the regulation of cell cycle progression through G1. Cyclin D1 is steroid-regulated (Refs. 17, 19, 38, and 39 and O. W. J. Prall, B. Sarcevic, E. A. Musgrove, C. K. W. Watts, and R. L. Sutherland, submitted), and recent data argue that activation of other G1 cyclins also contributes to steroid-induced mitogenesis (Ref. 39 and O. W. J. Prall, B. Sarcevic, E. A. Musgrove, C. K. W. Watts, and R. L. Sutherland, submitted). However, decreased cyclin D1 expression does not always accompany inhibition of breast cancer cell proliferation (18, 19). This manuscript further investigated the effects of antiprogestins and demonstrates that while regulation of cyclin/CDK function is likely to account for their inhibition of cell cycle progression, this is not mediated by regulation of cyclin abundance.

Accumulation of underphosphorylated pRB was coincident with the decrease in S phase fraction. Both were first evident at 9 h and reached a minimum after 18 h ORG 31710 treatment. These responses were preceded slightly by decreased cyclin D1- and cyclin E-associated kinase activity, which reached a minimum by 12 h. Since pRB is a key substrate for G1 cyclin-associated kinases, these data suggest that decreased pRB phosphorylation results from decreased CDK activity and that it is the presence of growth-inhibitory, underphosphorylated, pRB that ultimately mediates G1 arrest. Investigation of possible mechanisms underlying the decrease in CDK activity revealed that the abundance of the CDK inhibitor p21 was increased by 3- to 4-fold after 12–24 h treatment, when decreased kinase activity was observed. Antiprogestin induction of p21 mRNA was also observed, suggesting that the increased protein abundance was, at least in part, a result of either increased transcription or stabilization of p21 mRNA. Induction of p21 was not observed in either a progesterone receptor-negative cell line (MDA-MB-231) or a progesterone receptor-positive MCF-7 variant that is insensitive to antiprogestins. However, it was observed in BT 474 cells accompanying an antiprogestin-induced decrease in S phase fraction, indicating a close correspondence between p21 induction and growth inhibition. Regulation of the abundance of cyclin D1, Cdk4, Cdk6, or p27 did not appear to contribute to the decreased kinase activity, nor did the relative Cdk4 abundance in cyclin D1 immunoprecipitates alter after antiprogestin treatment (our unpublished data). The latter observation argues against the induction of p16INK4 or a related CDK inhibitor after antiprogestin treatment because this would be expected to displace Cdk4 from the complexes (24). Furthermore, preliminary examination of cyclin D1 immunoprecipitates from 35S-methionine-labeled T-47D cells did not provide evidence for alterations in complex composition other than increased p21 association after antiprogestin treatment (our unpublished data). Similarly, decreases in the abundance of cyclin E or Cdk2 sufficient to account for the decrease in cyclin E-associated kinase activity were not observed, but the relative abundance of p21 increased in cyclin E immunoprecipitates at times when kinase activity was reduced.

In a variety of cell types, ectopic expression of p21 leads to arrest in G1 phase (26, 40, 41), consistent with the hypothesis that induction of p21 might be responsible for the G1 phase arrest after antiprogestin treatment. Further experiments indicated that the observed increase in p21 abundance was likely to be sufficient to account for the decrease in cyclin D1-associated kinase activity. Although addition of p21 to recombinant cyclin D1/Cdk4 complexes inhibits kinase activity in a concentration-dependent fashion, the complexes retain near-maximal kinase activity in the presence of p21 levels representing 25–50% of the level required to completely inhibit kinase activity (42). Although the precise details of the p21 binding required for inhibition remain undefined, it is apparent that kinase activity is likely to decrease as the complexes approach saturation. In untreated T-47D cells a third of the total cellular p21 coimmunoprecipitated with cyclin D1 (Fig. 8Go), consistent with data from other cell types (e.g. Ref.43). This observation suggested that the cyclin D1 complexes contained p21 at a level near that required for inhibition of kinase activity. Addition of recombinant GST-p21 to cyclin D1 immunoprecipitates indicated that saturation of the cyclin D1 complexes with p21 would occur after an approximately 4-fold increase in bound p21 (Fig. 8Go) and thus that the observed ~3-fold increase in p21 coimmunoprecipitating with cyclin D1 was likely to be sufficient to decrease cyclin D1-associated kinase activity. The observation that the threshold for cyclin D1 function was increased after antiprogestin treatment (Fig. 9Go) is consistent with this interpretation since the abundance of CDK inhibitors is thought to set this threshold (24). Although the degree of induction of p21 in BT 474 was similar to that in T-47D, there was more Cdk4 and less p21 coimmunoprecipitated with cyclin D1 in BT 474, even after antiprogestin induction of p21 (Fig. 7CGo), suggesting a greater proportion of active complexes. This could then contribute to the modest level of the decrease in BT 474 S phase fraction after ORG 31710 treatment.

Because the activity of both cyclin D1 and cyclin E is required for progress into S phase (44, 45), inhibition of either could account for growth inhibition after antiprogestin treatment. However, despite the antiprogestin-induced decrease in both cyclin D1-associated and cyclin E-associated kinase activity, a 2.5-fold increase in cyclin D1 abundance alone prevented antiprogestin inhibition of cell cycle progression, apparently overriding effects on cyclin E-associated kinase activity. Thus, inhibition of cyclin D1-associated kinase activity appears to be a critical element in antiprogestin inhibition of cell cycle progression. An implication of this conclusion is that sensitivity to inhibition by antiprogestins may, in part, depend on the abundance of cyclin D1 and hence that the significant fraction, 30–50%, of breast cancers that overexpress cyclin D1 (31, 46, 47, 48, 49) may display altered sensitivity to antiprogestin therapy.

Occupancy of cyclin E complexes by p21 was not investigated in such detail as that of cyclin D1 complexes, but the correspondence between increased p21 binding and decreased kinase activity is consistent with increased p21 abundance contributing to the inhibition of cyclin E-associated kinase activity. However, it does not exclude other mechanisms, including alterations in the level or activity of the kinases and phosphatases controlling the level of Cdk2 phosphorylation and hence its activity. Furthermore, there is increasing evidence for growth-inhibitory effects of p21 not mediated via direct inhibition of CDK activity but rather by interaction with other cell cycle-regulatory pathways. For example, the activity of E2F, a transcription factor with a central role in cell proliferation, is repressed by p21 (50, 51). A possible mechanism is suggested by p21 disruption of the interaction between Cdk2, E2F, and the pRB-related proteins p107 and p130 (50, 51, 52), and this has been suggested to play a role in the inhibitory function of p21 (51). In addition, p21 binds the proliferating cell nuclear antigen (PCNA), blocking its ability to activate DNA polymerase {delta}, and the PCNA-binding domain alone is capable of blocking cell cycle progression (24). Since the PCNA-binding domain is a less effective growth inhibitor than the CDK-inhibitory domain (24), it is unlikely that inhibition of DNA replication by this mechanism makes a major contribution to antiprogestin inhibition of proliferation. Finally, p21 has recently been shown to inhibit the activity of kinases other than CDKs, i.e. the stress-activated protein kinases (also known as the c-Jun amino-terminal kinases) and protein kinase CK2 (53, 54), and it is conceivable that such inhibition also contributes to its growth-inhibitory effects.

The effects of antiprogestin on breast cancer cells are not limited to inhibition of cell proliferation. Other responses including induction of differentiation and apoptosis are apparent after several days’ antiprogestin treatment (55, 56). Although both differentiation and apoptosis can occur in the absence of p21 induction (24), increased p21 has been associated with differentiation in a number of cellular systems both in vivo and in vitro (24) and with retinoid induction of apoptosis in breast cancer cells (57). Overexpression of p21 in human melanocytes led to morphological changes characteristic of differentiation and increased melanin production, in some cases followed by cell death (41), while overexpression in MCF-7 and T-47D breast cancer cells led to apoptosis (58). These data suggest that p21 induction could contribute to antiprogestin effects other than growth arrest alone.

In summary, the data presented in this manuscript suggest a model for the effects of antiprogestin treatment on proliferation in which decreased activity of both cyclin D1/CDK and cyclin E/CDK and consequent decreased pRB phosphorylation result in inhibition of entry into S phase. Decreased kinase activity does not result from regulation of cyclin abundance but induction of the CDK inhibitor p21 accompanies these changes and is thus a likely mediator of the effects of antiprogestins on cell proliferation.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Cell Culture
The human breast cancer cells were obtained from the following sources: T-47D and MDA-MB-231, EG & G Mason Research Institute (Worcester, MA); MCF-7, Michigan Cancer Foundation (Detroit, MI); BT 474, American Type Culture Collection (Rockville, MD). T-47D {Delta}MTcycD1-3 are a clonal derivative of T-47D expressing ectopic cyclin D1 under the control of a metal-inducible truncated human metallothionein IIA promoter lacking steroid-responsive sequences (32, 35). RPMI 1640 medium was supplemented with 10 µg/ml human insulin (Actrapid, CSL-Novo, North Rocks, NSW, Australia), HEPES (20 mM), sodium bicarbonate (14 mM), and L-glutamine (6 mM). Stock cultures were maintained as previously described (36), in medium supplemented with 10% FCS and without antibiotics. Experiments used cells cultured in medium supplemented with 5% FCS. RU 486 (generously provided by Dr J-P Raynaud of Roussel-Uclaf, Romainville, France) and ORG 31710 (generously provided by Dr W. Schoonen, Organon, Oss, The Netherlands) were dissolved in ethanol at 1000- or 2000-fold final concentration and added to cells in exponential growth. Control cultures received ethanol to the same final concentration. Data presented are representative of at least two experiments. Cell cycle phase distribution was determined by flow cytometry (59).

Antiprogestin effects on cell number were measured using a colorimetric cell proliferation assay (CellTiter, Promega, Madison, WI). Cells (103) were seeded into 96-well plates, and the next day antiprogestin or vehicle was added to quadruplicate wells for each treatment. Plates were assayed at intervals and relative absorbances, i.e. relative cell numbers, were determined near the end of exponential growth for control cultures, after 6–7 days’ exposure to antiprogestin.

Recombinant p21
The coding region of p21 was amplified by PCR using pZL.WAF1 (60) as a template and p21N (TAC ATG GAT CCA TGT CAG AAC CGG CTG GGG A) and p21C (AGA CTG AAT TCT TAG GGC TTC CTC TTG GAG A) as primers. The resulting fragment was cloned into the BamHI/EcoRI sites of the expression vector pGEX-2t (Pharmacia, Uppsala, Sweden) to yield pGEX-p21. To prepare GST-p21, Eschericia coli were transformed with pGEX-p21, and expression of the protein was induced by incubation with 0.1 mM isopropylthioglycoside) (2.5 h, room temperature). Frozen cell pellets were lysed by sonication at 4 C after resuspension in PBS with protease inhibitors (1 mM phenylmethylsulfonylfluoride, 0.5 M EDTA, 0.05% (vol/vol) ß-mercaptoethanol, 10 µg/µl aprotonin, 10 µg/µl leupeptin). After addition of 0.5% Triton X-100, the lysate was centifuged at 4 C. The supernatant was then incubated with gentle rotation at 4 C for 1 h with 0.5 ml of a 50% (vol/vol) suspension of glutathione-agarose (Sigma, St.Louis, MO). The resin was washed once with ice-cold PBS/0.05% ß-mercaptoethanol/0.5% Triton and twice with ice-cold PBS/0.05% ß-mercaptoethanol and finally resuspended in 0.5 ml ice-cold PBS/0.05% ß-mercaptoethanol/0.1% azide. The fusion protein was then eluted with 50 mM Tris-Cl, pH 8.0, 15 mM reduced glutathionine, 0.05% ß-mercaptoethanol, and the purity of fusion protein was assessed by PAGE followed by Coomassie blue staining.

Cell Lysis, Western Blot Analysis, and Immunoprecipitation
Cells were lysed either as described below for cyclin D1-associated kinase assays or as previously described, using lysis buffer consisting of 50 mM HEPES (pH 7.5), 150 mM NaCl, 10% (vol/vol) glycerol, 1% Triton X-100, 1.5 mM MgCl2, 1 mM EGTA, 10 µg/ml aprotinin, 10 µg/ml leupeptin, 1 mM PMSF, 200 µM sodium orthovanadate, 10 mM sodium pyrophosphate, and 100 mM NaF (32, 61). Similar results were obtained from Western blotting or immunoprecipitation using either lysis technique.

Cell lysates were precleared by incubation with protein A-Sepharose beads (Zymed, San Francisco, CA) (1 h, 4 C) then immunoprecipitated by incubation (3 h, 4 C) with protein A-Sepharose beads that had been conjugated with either an anti-cyclin E antibody (C-19, Santa Cruz Biotechnology, Santa Cruz, CA) or rabbit polyclonal anti-cyclin D1 serum [raised against a human cyclin D1-GST fusion protein (35)]. In some experiments the antibodies were chemically cross-linked to the protein A-Sepharose beads by incubation in 5 mg/ml dimethyl pimelimidate/0.2 M sodium tetraborate (pH 9.0) for 30 min at room temperature, essentially as described (62). The immunoprecipitated proteins were then washed as previously described (35). In the experiments presented in Fig. 8Go, recombinant GST-p21 was added to the immunoprecipitates, and the samples were incubated at 30 C for 1 h with vortexing every 10 min. The beads were then washed with 50 mM HEPES (pH 7.5), 1 mM dithiothreitol before resuspension in SDS-PAGE sample buffer.

Samples of immunoprecipitated or total protein in SDS-PAGE sample buffer were heated to 95 C for 3 min, then separated by SDS-PAGE and transferred to nitrocellulose. Specific proteins were visualized by chemiluminescence (Dupont NEN, Boston, MA) after incubation (2–4 h at room temperature or overnight at 4 C) with the following primary antibodies: p21 antiserum kindly provided by Dr David Beach (Cold Spring Harbor, NY); cyclin E (HE12), Cdk2 (M2), Cdk4 (C-22), and Cdk6 (C-21) antibodies from Santa Cruz Biotechnology; cyclin D1 antibody (DCS6) from Novocastra, Newcastle-upon-Tyne, U.K.; pRB (14001A) antibodies from Pharmingen (San Diego, CA); p21 (C24420) and p27 (K25020) antibodies from Transduction Laboratories (Lexington, KY). Relative abundance was quantitated using a Molecular Dynamics (Sunnyvale, CA) densitometer and IP LabGel analysis software (Signal Analytics, Vienna, VA).

Kinase Assays
The histone H1 kinase activity of cyclin E immunoprecipitates was measured as previously described for Cdk2 assays (35) using 10 µg histone H1 as substrate. For cyclin D1-associated kinase assays, cells were harvested and lysed as previously described using kinase lysis buffer [50 mM HEPES (pH 7.5), 1 mM dithiothreitol, 150 mM NaCl, 1 mM EDTA, 2.5 mM EGTA, 0.1% Tween-20, 10% glycerol, 10 mM ß-glycerophosphate, 1 mM NaF, 0.1 mM sodium orthovanadate, 10 µg/ml leupeptin, 10 µg/ml aprotinin, and 0.1 mM PMSF] (35, 63). Kinase activity of cyclin D1 immunoprecipitates of these lysates was measured using either a pRB(379–928)-maltose binding protein fusion protein, GST-pRB(769–921) fusion protein substrate (Santa Cruz) or GST-pRB(773–928) (64) as previously described (35). After termination of kinase reactions, samples were incubated at 90 C for 2 min in SDS sample buffer and separated using 10% SDS-PAGE. Relative intensities were quantitated using a Molecular Dynamics PhosphorImager Scanner (model 445 SI) followed by analysis using IP LabGel analysis software (Signal Analytics) or in some cases after exposure to x-ray film as decribed above for Western analysis. The degree of background phosphorylation in cyclin D1-associated kinase assays was estimated from parallel control samples either immunoprecipitated using preimmune serum or assayed after incubation of immunoprecipitates with an excess of GST-p21 (30 min, 30 C) and has been subtracted in the data presented in Fig. 3AGo.

RNA Isolation and Northern Analysis
Total RNA was extracted (using a guanidinium isothiocyanate-cesium chloride procedure) and blotted as previously described, using 20 µg total RNA/lane (46). The membranes were hybridized overnight at 50 C in 50% (vol/vol) formamide, 2x SSPE (0.3 M NaCl, 20 mM NaH2PO4, 2 mM EDTA, pH 7.4), 1% (wt/vol) SDS, 0.5% (wt/vol) low fat skim milk (Diploma, St Kilda, Victoria, Australia), 10% (wt/vol) dextran sulfate (Mr 500,000), 200 µg/ml yeast RNA, 40 µg/ml polyadenylic acid (5'), 500 µg/ml salmon sperm DNA. The 2.1-kb p21 cDNA from pZL.WAF1 (60) was labeled with [{alpha}-32P]dCTP (Amersham Australia, North Ryde, New South Wales, Australia; specific activity ~3000 Ci/mmol) to a specific activity of approximately 1 x 109 cpm/µg DNA using the Multiprime DNA labeling kit (Amersham Australia) then added to the hybridization mix at a final concentration of >= 10 ng/ml. The membranes were washed at a highest stringency of 0.2 x SSC (30 mM NaCl, 3 mM sodium citrate, pH 7.0), 1% SDS at 65 C and exposed to Kodak X-OMAT film at -70 C. Equivalent RNA loading was verified as previously described (46) by hybridizing membranes with a [{gamma}-32P]ATP end-labeled oligonucleotide complementary to 18S rRNA.

Statistical Analysis
Data pooled from multiple experiments were analyzed using Statview II software (Abacus Concepts, Inc, Berkeley, CA). The significance of differences from control was determined using a one-tailed t test.


    ACKNOWLEDGMENTS
 
The authors thank Drs David Beach and Jiri Lukas for supplying antibodies.


    FOOTNOTES
 
Address requests for reprints to: Elizabeth A. Musgrove, Cancer Research Program, Garvan Institute of Medical Research, St. Vincent’s Hospital, Sydney, New South Wales 2010 Australia.

This study was supported by research grants from the National Health and Medical Research Council of Australia and the New South Wales State Cancer Council.

Received for publication September 16, 1996. Accepted for publication October 3, 1996.


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