Cisplatin Potentiates 1,25-Dihydroxyvitamin D₃-induced Apoptosis in Association with Increased Mitogen-activated Protein Kinase Kinase Kinase 1 (MEKK-1) Expression ¹

In addition to its classical effects on bone and mineral metabolism, calcitriol, or 1,25D₃,⁴ has antiproliferative activity against a variety of solid tumors including those of the breast, prostate, and colon (1–5). 1,25D₃ also inhibits the in vitro and in vivo growth of the murine SCC cell line SCCVII/SF (6); antiproliferative activity in this model is associated with G₀-G₁ cell cycle arrest (7, 8) and induction of apoptosis (9). 1,25D₃ and 1,25D₃ analogues have been studied in combination with several cytotoxic drugs including platinum-containing agents (7, 10, 11), taxanes (12–14), and DNA-intercalating agents (13, 15). Such combinations display antitumor activity greater than that observed for any single agent. The combination of 1,25D₃ with cisplatin demonstrates increased activity against breast (11), prostate (10), and SCC cancer cell lines (7). Optimal cytotoxicity for the combination in SCC requires pretreatment of cells with 1,25D₃ for at least 24 h before treatment with cisplatin (7). The molecular mechanism by which antitumor activity is enhanced for this combination is unknown.

Cisplatin cytotoxicity results from the formation of bifunctional, intrastrand DNA adducts (16). In eukaryotic cells, DNA damage results in stabilization and activation of the tumor suppressor p53 (17). Such activation results in increased transcription of p53 target genes including the cyclin-dependent kinase inhibitor p21^Waf1/Cip1 (18), the proapoptotic Bcl-2 family member Bax (19), and other genes implicated in cell cycle arrest, DNA repair, and apoptosis (20). p53 signaling plays a role in dictating the outcome of cisplatin exposure; modulation of the expression of p53 or p53 target genes alters cellular sensitivity to cisplatin (21–26). In addition, cisplatin cytotoxicity may be attenuated by factors that regulate the intracellular accumulation of platinum, DNA repair, and tolerance to DNA damage (27).

Stress and survival signaling molecules also play a critical role in determining sensitivity to cytotoxic agents such as cisplatin. For example, in certain cancer cell lines, genotoxic agents activate MEK-Erk signaling, resulting in decreased cytotoxicity of those agents (reviewed in Ref. 28). Ovarian carcinoma cell lines demonstrate Erk activation and reduced cytotoxicity upon exposure to cisplatin, and treatment of these cells with the MEK inhibitor PD98059 increases cisplatin cytotoxicity (29, 30). Several reports have also demonstrated an important role for stress signaling molecules such as MEKK-1 in regulating cisplatin sensitivity. MEKK-1 becomes phosphorylated and is subsequently cleaved by caspase-3 upon treatment with cisplatin (31). Cleavage leads to the loss of intact MEKK-1 and the generation of a 91-kDa constitutively active COOH-terminal MEKK-1 catalytic domain that is released into the cytoplasm (31). Aberrant expression of unregulated MEKK-1 activity in the cytoplasm leads to the generation of proapoptotic signals via the c-Jun N-terminal kinase kinase (SEK1)-JNK-Jun and MKK3/6-p38 MAPK stress pathways and ultimately leads to apoptosis (32–34).

Recently, we reported that the proapoptotic effects of 1,25D₃ in SCC are associated with up-regulation of MEKK-1 before the onset of apoptosis (9). In addition, 1,25D₃ induces caspase-dependent proteolytic cleavage of MEK with consequent inhibition of MEK-Erk mitogenic/survival signaling. 1,25D₃ also decreases the level of phosphorylated/activated c-Jun, and either maintains or induces phospho-p38 MAPK (9). Based on these findings, we proposed that the loss of MEK/Erk survival signaling and the increase in MEKK-1/p38 MAPK stress signaling shift the balance from cell survival toward cell death in response to 1,25D₃ treatment. Interestingly, MEK cleavage and MEKK-1 up-regulation appear to be selective markers for apoptosis induced by 1,25D₃ [i.e., they are either not observed or observed to a lesser extent for apoptosis induced by traditional cytotoxic agents, including cisplatin (9)]. In fact, in SCC cells, 1,25D₃ and cisplatin display opposite effects on the levels of intact MEKK-1 and phospho-Jun (9).

We have focused on alterations in cellular platinum, formation of platinum-DNA adducts, and effects on known components of cisplatin and 1,25D₃ signaling pathways in an attempt to elucidate the molecular mechanism of enhanced antitumor activity for the combination of 1,25D₃ plus cisplatin. Here, we report that SCC cells treated with the combination of 1,25D₃ and cisplatin exhibit an apoptotic response that is characteristic of 1,25D₃ and that the up-regulation of MEKK-1 observed for 1,25D₃ alone is potentiated by cotreatment with cisplatin. These data suggest that 1,25D₃ pretreatment commits cells to undergo apoptosis using a specific molecular pathway and that traverse along this pathway is increased when cells receive an additional, low-level genotoxic stimulus.

1,25D₃ (Hoffmann-LaRoche, Nutley, NJ) was reconstituted in 100% EtOH and stored, protected from light, under nitrogen at −70°C. All handling of 1,25D₃ was performed with indirect lighting. 1,25D₃ was diluted in tissue culture medium just before use. Cisplatin (Platinol-AQ; Bristol Laboratories, Princeton, NJ) was obtained as a 1 mg/ml solution and diluted in sterile saline or tissue culture medium immediately before use.

Murine SCC cells (SCCVII/SF) were obtained and passaged as described previously (6). Cells were plated in RPMI 1640 supplemented with 12% fetal bovine serum (HyClone Laboratories, Logan, UT) and 1% penicillin/streptomycin sulfate and incubated at 37°C in a humidified atmosphere containing 5% CO₂. Cells were allowed to attach overnight and then treated and analyzed as described below. EtOH, used as a solvent control, never exceeded a final concentration of 0.0004%.

SCC cells (1.5 × 10⁴ cells/ml) were seeded in a volume of 0.1 ml into 96-well tissue culture plates. Cells were pretreated for 24 h by adding medium alone or medium containing 1,25D₃ directly to the wells. Cells were either incubated without further treatment or treated with medium containing cisplatin for an additional 24 h. The final volume in each well was 0.2 ml. To quantitate cell viability, the cells were incubated with 20 μl of 0.5% MTT for 90 min at 37°C. The medium was removed, and the cells were solubilized for 20 min in 10% SDS/10 mm HCl. Absorbance was read with an ELISA plate reader (Bio-Tek Instruments, Winooski, VT) at a wavelength of 490 nm. Median doses for each drug were determined from the dose-response data using CalcuSyn software (Biosoft, Ferguson, MO). Drug interactions were quantitated using the equation CI = (D)₁/(Dx)₁ + (D)₂/(Dx)₂. (D)₁ and (D)₂ are the doses of drug 1 and drug 2 that, when given in combination, inhibit cell growth by a specified percentage. (Dx)₁ and (Dx)₂ are the doses of drug 1 and drug 2 that, when given individually, inhibit cell growth by the same percentage. CI values of <1, 1, and >1 indicate synergism, additivity, and antagonism between the drugs, respectively.

SCC cells (2.5 × 10⁶ cells/T75 flask) were treated with either EtOH or 10 nm 1,25D₃. After 24 h, the medium was replaced with fresh medium containing cisplatin (0.5 μg/ml), and cells were incubated for an additional 0.5–2.0 h at 37°C. At the conclusion of the incubation, the tissue culture flasks were placed on ice, and the cells were harvested by rapid mixing with glass beads. The cells were collected by centrifugation (5 min at 200 × g at 4°C) and washed in ice-cold PBS. The resulting cell pellets were resuspended in ice-cold PBS, and an aliquot of each cell suspension was removed for cell count determination. The remaining cells were transferred to an Eppendorf tube and collected by centrifugation. The final cell pellet was dissolved in 0.25% Triton X-100, and total cellular platinum was determined by flameless atomic absorption spectroscopy using a Perkin-Elmer model 1100 spectrometer (Perkin-Elmer, Norwalk, CT) monitoring 265.9 nm, as described previously (35). Platinum concentration was determined by comparison with a standard curve and was expressed on a per cell basis.

Triplicate sets of three T75 flasks each of SCC cells (1.25 × 10⁶ cells/flask) were pretreated with either EtOH or 10 nm 1,25D₃ for 24 h. Cisplatin was added directly to pretreated cultures to a final concentration of 0.5 μg/ml, and cells were incubated for an additional 2 h. The tissue culture medium was removed, and adherent cells were recovered by trypsinization. DNA extraction was initiated by pooling cells from three identically treated flasks into a single tube. Cells were collected by centrifugation and washed twice with PBS. Cell pellets were lysed in a buffer containing 10 mm Tris-Cl (pH 8.0), 0.1 m NaCl, 0.1 mm EDTA, 0.5% SDS, and 20 μg/ml RNase and incubated for 5 h at 37°C. Proteinase K was then added to a final concentration of 100 μg/ml, and the preparation was incubated overnight at 50°C. DNA was extracted with phenol-chloroform and precipitated with EtOH. DNA precipitates were suspended in distilled H₂O, and the DNA yield was determined by reading the absorbance at 260 nm. Platinum-DNA adduct levels were quantitated using a ³²P postlabeling assay (36) and are expressed as fmol adducts/μg DNA.

SCC cells (either 2 × 10⁵ or 4 × 10⁵ cells/T75 flask) were pretreated for 24 h with EtOH or 10 nm 1,25D₃. Cells were incubated without further treatment, or cisplatin was added directly to tissue culture medium to a final concentration of 0.5 or 5.0 μg/ml. Cells were incubated for an additional 24 h. For caspase-3 Western blots only, adherent cells were scraped up into the medium containing floating cells. For all other analyses, medium in the flasks was swirled and removed to collect floating cells. Attached cells were then scraped off the flasks into PBS and placed in a separate tube. Protein extracts were obtained by lysing the pooled, attached, or floating cell populations in Triton X-100/SDS buffer as described previously (9). Equivalent amounts of protein from each sample were electrophoresed under denaturing conditions on SDS-polyacrylamide gels. Membranes were blocked and probed as described previously (8, 9). The antibodies used were anti-caspase 3 (210-807-C100; Alexis Corp., San Diego, CA), anti-PARP (C2.10; Enzyme Systems Products, Livermore, CA), anti-p53 (sc-99; Santa Cruz Biotechnology, Santa Cruz, CA), anti-Bax (13686E; PharMingen, San Diego, CA), anti-p21^Cip1 (13436E; PharMingen), anti-phospho-MEK1/2 (9121S; Cell Signaling Technologies, Beverly, MA), anti-phospho-Erk1/2 (sc-7383; Santa Cruz Biotechnology), anti-phospho-c-Jun (sc-822; Santa Cruz Biotechnology), and anti-MEKK-1 (sc-252; Santa Cruz Biotechnology). Secondary antimouse and antirabbit horseradish peroxidase-conjugated antibodies were from Amersham Biosciences (Piscataway, NJ). Actin was detected using the actin (Ab-1) kit from Oncogene Research Products (Boston, MA). Caspase-3 antibody reactions were visualized using SuperSignal substrate (Pierce, Rockford, IL). All other reactions were visualized using Renaissance Enhanced Chemiluminescence Reagents (NEN Life Science Products, Boston, MA).

To explore the nature of the interaction between 1,25D₃ and cisplatin in SCC, cells were treated with varying concentrations of either 1,25D₃ or cisplatin alone, or they were pretreated with 1,25D₃ followed by cisplatin (1,25D₃/cisplatin); antiproliferative effects were measured using the MTT assay. As shown in Fig. 1A, greater inhibition of cell growth was observed for the combination than for either single agent. From these dose-effect data, it was determined that the concentrations of 1,25D₃ and cisplatin required for 50% growth inhibition were 3–8-fold lower, respectively, when the drugs were used in combination than when used individually (Fig. 1A). These data were also used to determine CI values as described in “Materials and Methods.” For most of the dose combinations tested, a CI value of <1 was obtained, indicating that the interaction between 1,25D₃ and cisplatin is synergistic (Fig. 1B). Median dose-effect analysis also revealed apparent antagonism between the drugs when low concentrations of 1,25D₃ were used in combination with higher cisplatin concentrations. However, no such antagonism is observed using in vitro clonogenic, in vivo excision clonogenic, or tumor outgrowth assays (7).

To dissect the mechanism for the enhanced antitumor effect of 1,25D₃/cisplatin therapy, a single dose of each agent was selected for combination, and the effects on induction of apoptosis were determined by monitoring caspase-3 cleavage. SCC cells were pretreated with either vehicle (EtOH) or 10 nm 1,25D₃. At this concentration, 1,25D₃ treatment results in 50% growth inhibition (Fig. 1A). Cisplatin was added directly to pretreated cultures to a final concentration of 0.5 μg/ml. This concentration of cisplatin was anticipated to interact favorably with 1,25D₃, based on the results presented in Fig. 1. 1,25D₃ induced apoptosis, as evidenced by the accumulation of caspase-3 cleavage fragments (Fig. 2). This agrees with our previous data demonstrating cleavage of the caspase-3 substrate, PARP, in 1,25D₃-treated SCC (9). Caspase-3 cleavage was not detected in cells treated with cisplatin alone (Fig. 2). However, in cells treated with 1,25D₃/cisplatin, caspase-3 cleavage was consistently enhanced above that observed for cells treated with 1,25D₃ alone (Fig. 2, A and B). These data suggest that the increase in antiproliferative activity observed for the combination of 1,25D₃ plus cisplatin is attributed, at least in part, to an increase in apoptosis.

To explore whether 1,25D₃ increases cisplatin toxicity in SCC by increasing either cellular platinum accumulation or the formation of DNA adducts, SCC cells were pretreated for 24 h with vehicle or 1,25D₃ and then treated with 0.5 μg/ml cisplatin for specified times. These times were selected based on observations that a 1-h exposure to cisplatin was sufficient to observe an increase in DNA platination in HL60 cells induced to differentiate with 1,25D₃ (37). Cells were harvested, and we measured either cellular platinum concentration (Table 1) or platinum-DNA adducts (Table 2). Cellular platinum concentrations were not statistically different in cells treated with 1,25D₃/cisplatin as compared with cells treated with cisplatin alone at any of the time points examined. Similarly, platinum-GG adducts, the major cross-linked species in cisplatin-treated cells (38), and platinum-AG adducts were not statistically different in cells treated with cisplatin alone compared with cells treated with 1,25D₃/cisplatin.

We subsequently tested the hypothesis that the increased apoptosis observed in SCC cultures treated with 1,25D₃/cisplatin results from a 1,25D₃-mediated increase in cisplatin signaling. After treatment with 1,25D₃, cisplatin, or 1,25D₃/cisplatin, two distinct populations of cells were evident; one adherent and the other floating (no floating cells were present in control cultures). Our prior work with 1,25D₃ demonstrated different molecular changes in these two populations (9); therefore, they were harvested and analyzed separately. Because the concentration of cisplatin used in these studies (0.5 μg/ml) did not induce caspase-3 activation (Fig. 2), in some experiments, cells were also treated with a 10-fold higher concentration of cisplatin as a positive control for cisplatin-induced apoptosis. Whole cell extracts were prepared, and immunoblot analysis was carried out for p53, p21, and Bax (to monitor p53 signaling) and phospho-c-Jun (to monitor JNK activation). Cleavage of PARP was also assessed so that molecular changes could be correlated with induction of programmed death.

Treatment with 0.5 μg/ml cisplatin modestly increased levels of full-length PARP in adherent cells and had little affect on PARP levels in floating cells (Fig. 3A). Based on the lack of PARP and caspase-3 cleavage (Figs. 2A and 3A) and the absence of membrane blebbing (data not shown), we conclude that this concentration of cisplatin, although promoting rounding and detachment of cells, did not induce apoptosis. However, SCC cells possess the molecular machinery to undergo cisplatin-induced apoptosis because treatment with 5.0 μg/ml cisplatin induced partial PARP cleavage in adherent cells and complete PARP cleavage in floating cells (Fig. 3A). 1,25D₃ alone induced a significant decrease in PARP expression in the attached cells and a complete loss in the floating cells (Fig. 3A). In addition, 1,25D₃ induced pronounced membrane blebbing in the floating cell population. Thus, in contrast to 0.5 μg/ml cisplatin, the population of floating cells in 1,25D₃-treated cultures is apoptotic. This is in agreement with our previous findings (9). When 1,25D₃ and 0.5 μg/ml cisplatin were combined, modulation of PARP levels resembled the effect of treatment with 1,25D₃ alone. However, when 1,25D₃ and 5.0 μg/ml cisplatin were combined, modulation of PARP levels was more similar to the effect of high-dose cisplatin alone (Fig. 3A).

We next assessed the expression of p53 and its downstream transcriptional targets, p21^Waf1/Cip1 and Bax, under the various treatment conditions. Treatment of cells with cisplatin (0.5 μg/ml) alone weakly induced p53 and its targets in both cell populations (Fig. 3, A and B). p53 induction was dose dependent, with much stronger induction observed at high (5.0 μg/ml) cisplatin concentrations (Fig. 3A). In contrast, 1,25D₃ reduced p53, p21, and Bax expression in the adherent cells and p53 and p21 expression in the floating cells (Fig. 3, A and B). It is intriguing to note that although Bax (21 kDa) was reduced in adherent cells after 1,25D₃ treatment, it was maintained at control levels in floating cells (Fig. 3B and discussed below). In addition, an 18-kDa Bax cleavage fragment was also detected in the floating cells (data not shown). Analogous to their differential effects on p53 expression, cisplatin and 1,25D₃ treatment also led to differential effects on the stress signaling kinase, JNK. Whereas cisplatin treatment resulted in elevated c-Jun phosphorylation, phosphorylation was decreased to below control levels after 1,25D₃ treatment (Fig. 3B). For each of the proteins examined, the molecular profile observed for cells treated with 1,25D₃/cisplatin (0.5 μg/ml) resembled that of cells treated with 1,25D₃ alone, whereas the profile observed for cells treated with 1,25D₃/cisplatin (5.0 μg/ml) resembled that of cells treated with cisplatin 5.0 μg/ml) alone ((Fig. 3, A and B).

The inhibitory effect of 1,25D₃ on cisplatin signaling makes it unlikely that the increase in antitumor activity for 1,25D₃/cisplatin is due to 1,25D₃ enhancement of cisplatin cytotoxicity. We therefore explored the alternative hypothesis that cisplatin enhances 1,25D₃ proapoptotic signaling in SCC cells. In accord with our previous report (9), the apoptotic, floating cell population from 1,25D₃-treated cultures demonstrated strongly reduced levels of MEK expression associated with the appearance of a 33-kDa cleavage fragment (Fig. 4A). As anticipated, the 1,25D₃-mediated reduction in MEK resulted in reduced phosphorylation of Erk1/2, its downstream target (Fig. 4A; Ref. 9). In contrast, cisplatin treatment resulted in increased Erk phosphorylation/activation in the adherent cells (although MEK expression did not change; Fig. 4A). In general, the MEK/Erk profile in 1,25D₃/cisplatin-treated cells resembled that of cells treated with 1,25D₃ alone, except that the 33-kDa MEK cleavage fragment was further increased in the floating cells after treatment with 1,25D₃/cisplatin as compared with cells treated with 1,25D₃ alone (Fig. 4A).

When administered as a single agent, cisplatin (0.5 μg/ml) caused little or no increase in MEKK-1 expression in the adherent cell population and reduced expression in the floating cells (Fig. 4B). As we reported previously, 1,25D₃ treatment was associated with up-regulation of the proapoptotic stress signaling kinase MEKK-1 in both adherent and floating cells (Fig. 4B; Ref. 9). MEKK-1 expression was increased to a similar extent in the cells that remained adherent after treatment with 1,25D₃ alone and 1,25D₃/cisplatin (Fig. 4B). After treatment with 1,25D₃/cisplatin, a further increase in MEKK-1 expression was observed in the floating, apoptotic cells, which exceeded that observed for 1,25D₃ alone (Fig. 4B). These data suggest that cisplatin enhances 1,25D₃ proapoptotic signaling through MEKK-1.

Antitumor activity is enhanced for combinations of 1,25D₃ (or 1,25D₃ analogues) with cytotoxic drugs including platinum-containing agents (7, 10, 11) and taxanes (12–14). The clinical implications of these laboratory findings are as follows: (a) for a fixed dose of cytotoxic drug, addition of 1,25D₃ may improve the antitumor response observed; and (b) a cytotoxic drug may be given at a reduced and potentially less toxic dose in combination with 1,25D₃ without loss of activity. As a step toward clinical development of 1,25D₃/cytotoxic combinations, Phase I clinical trials of 1,25D₃ plus carboplatin or paclitaxel are currently in progress at our institution (39). However, the mechanism by which antitumor activity is increased for such combinations is unclear. We report here that enhancement of antitumor activity for the combination of 1,25D₃ plus cisplatin in SCC is associated with increased apoptosis. By surveying known components of the 1,25D₃ and cisplatin signaling pathways, we find that the increase in apoptosis for the combination is associated with up-regulation of MEKK-1 to levels that exceed those observed for either agent alone. Our analysis of other molecular markers that are selectively induced by either 1,25D₃ or cisplatin reveals that the molecular profile of cells treated with 1,25D₃/cisplatin most closely resembles that of 1,25D₃. The mechanistic implication of these results is that cisplatin potentiates 1,25D₃ proapoptotic activity in SCC.

The ability of MEKK-1 to stimulate apoptosis is well documented (31, 34). These studies demonstrate that MEKK1-mediated apoptosis depends upon proteolytic processing at a caspase-3 cleavage site. Such processing results in a COOH-terminal fragment of MEKK-1 (ΔMEKK1) that stimulates apoptosis. We reported previously (9) that 1,25D₃ alone induces MEKK-1 up-regulation in both adherent and floating cell populations and that NH₂-terminal proteolytic processing of MEKK-1 (distinct from caspase-3 cleavage) occurs in those cells that become apoptotic after treatment. Ectopic overexpression of MEKK-1 has been reported to lead to accumulation of MEKK-1 in the cytosol and subsequent apoptosis (31). These findings have led to the concept that aberrant expression of MEKK-1 in the cytosol triggers the generation of apoptotic signals. Thus, if the strong up-modulation of MEKK-1 observed in 1,25D₃-treated cells results in its cytosolic expression, this may “prime” the cells to initiate an apoptotic signal upon the addition of a low-level cytotoxic stimulus. This hypothesis is supported by previous findings of Widmann et al. (40), who demonstrated that acute expression of ΔMEKK1 in 3T3 cells 17 h before cisplatin results in greater inhibition of cell number than treatment with cisplatin alone.

Based on our findings, we propose that the up-regulation of MEKK-1 resulting from 1,25D₃ pretreatment in SCC results in apoptotic sensitization, and this accounts for the increase in apoptosis observed for the combination of 1,25D₃ and cisplatin. However, it appears that cisplatin triggers a greater apoptotic response using a mechanism that is more characteristic of 1,25D₃ than cisplatin. Thus, one possible interpretation of our data is that 1,25D₃ pretreatment and consequent MEKK-1 up-regulation commit cells to a particular apoptotic pathway and that traverse along this pathway is enhanced in the presence of cisplatin (see Fig. 5). The signals downstream of MEKK-1 that lead to apoptosis remain largely undefined. However, one recent study demonstrates that conformational changes in the proapoptotic Bcl-2 family member Bak occur in response to MEKK-1 activation in cisplatin-treated melanoma cell lines (41). Future studies will be required to explore the exact biological consequences of 1,25D₃-induced MEKK-1 up-regulation and its effects on Bak conformation in SCC.

The molecular survey done here indicates 1,25D₃ modulates multiple factors that could increase cellular susceptibility to apoptosis. Of note in this regard are the changes we observe in the proapoptotic Bax protein, which promotes cell death by modulating mitochondrial release of proapoptotic factors such as cytochrome c (42). We find that there is an association between Bax expression and apoptosis induction in 1,25D₃-treated SCC such that although Bax expression is reduced in viable cells that remain adherent, it is retained in the floating, apoptotic cells. In addition, Bax is cleaved to an 18-kDa species in the apoptotic cell population (data not shown). An 18-kDa calpain cleavage fragment of Bax has been identified (43) and reported to possess proapoptotic activity (44, 45), making it seem likely that this species also retains proapoptotic function in SCC cells. A role for 1,25D₃-mediated alterations in Bax subcellular distribution and cleavage in the proapoptotic effects of 1,25D₃ has been reported previously in MCF-7 breast cancer cells (46). It is interesting to note that the effects of 1,25D₃ on Bax expression appear to be p53 independent because Bax is retained in floating cells with reduced p53 expression (Fig. 3, A and B).

We reported previously that 1,25D₃-mediated apoptosis is associated with caspase-dependent cleavage of MEK, resulting in down-regulation of survival signaling through Erk (9). MEK cleavage, as evidenced by loss of full-length MEK and the appearance of a 33-kDa cleavage fragment, was consistently observed in the studies reported here (Fig. 4A). As compared with treatment with 1,25D₃ alone, MEK cleavage fragments were further increased in cells that became apoptotic after treatment with 1,25D₃/cisplatin. Cisplatin alone did not induce MEK cleavage or inhibition of phospho-Erk1/2 in the floating cells and did not induce apoptosis as evidenced by a lack of caspase-3 and PARP cleavage (Figs. 2A and 3A). Thus, these data indicate that cisplatin further potentiates the 1,25D₃-mediated loss of survival signaling and are consistent with our hypothesis that cisplatin increases signaling through the 1,25D₃ proapoptotic pathway.

Treatment with cisplatin alone has been reported to induce Erk phosphorylation/activation in ovarian cancer cell lines (47). In these cells, treatment with cisplatin in the presence of the MEK inhibitor PD98059 results in a reduction in cisplatin-induced p53 accumulation (47). These studies suggest Erk is one of the regulators of p53 protein accumulation during the DNA damage response to cisplatin. It is therefore tempting to speculate that the 1,25D₃-mediated reduction in MEK/Erk signaling may, in part, account for the reduction in p53 observed in cells treated with 1,25D₃ alone and the block to p53 induction observed in cells treated with 1,25D₃ plus cisplatin (Fig. 3A). The inhibitory effect of 1,25D₃ on p53 induction has been reported previously; pretreatment of MCF-7 cells with the 1,25D₃ analogue EB1089 enhanced Adriamycin cytotoxicity and blocked Adriamycin induction of p53 (48).

p21^Waf1/Cip1 is transcriptionally up-regulated by p53 (18). Thus, 1,25D₃-mediated inhibition of p53 may explain the suppression of p21 observed in 1,25D₃-treated SCC cells (Fig. 3A). After DNA damage, p21 promotes cell cycle arrest by interacting with and inhibiting both cyclin-cyclin-dependent kinase complexes (49) and PCNA (50, 51). Although the interaction between p21 and PCNA results a block in DNA replication, this association may actually be required for efficient removal of DNA adducts via the nucleotide excision repair pathway. Consistent with this hypothesis, p21-deficient HCT116 colon cancer cells show a reduced capacity to repair cisplatin-damaged DNA, and this nucleotide excision repair deficiency is overcome by reintroduction of wild-type p21 but not p21 mutants lacking a PCNA-binding domain (52). More recent studies suggest that PCNA remains associated with DNA for prolonged periods in p21-deficient fibroblasts, which may ultimately result in reduced PCNA recycling and accumulation of unrepaired lesions (53). Although we did not observe a difference in DNA adduct formation after a 2-h exposure to cisplatin (Table 2), it is possible that 1,25D₃, by reducing p21 expression, may inhibit the repair of cisplatin-induced DNA adducts at later time points and thereby contribute to cytotoxicity. Studies to test this hypothesis are currently in progress.

Finally, we show that for most dose combinations tested, there is a synergistic interaction between 1,25D₃ and cisplatin in SCC (Fig. 1B). Similar observations have been reported for both breast and prostate cancer cells (10, 11). Our studies in SCC also reveal apparent antagonism between these agents, an effect that is restricted to combinations using very low concentrations of 1,25D₃ and concentrations of cisplatin ≥ 0.225 μg/ml. The short half-life of cisplatin in plasma as compared with tissue culture medium makes extrapolation of our in vitro cisplatin exposures to in vivo exposures difficult. However, we consistently find that the 1,25D₃/cisplatin combination exhibits greater antitumor activity than either single agent in in vivo excision clonogenic assays and tumor outgrowth studies (7).⁵ These in vivo data predict enhanced rather than diminished clinical activity for this combination.

We thank Robert Reuger for maintenance of SCC cultures and excellent technical assistance. We also thank members of the University of Pittsburgh Division of Hematology-Oncology writing group for their careful review of the manuscript.