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. Author manuscript; available in PMC: 2014 Oct 1.
Published in final edited form as: Int J Cancer. 2010 Aug 15;127(4):977–988. doi: 10.1002/ijc.25112

Overexpression of DDB2 enhances the sensitivity of human ovarian cancer cells to cisplatin by augmenting cellular apoptosis

Bassant M Barakat 1, Qi-En Wang 1, Chunhua Han 1, Keisha Milum 1, De-Tao Yin 1, Qun Zhao 1, Gulzar Wani 1, El-Shaimaa A Arafa 1, Mohamed A El-Mahdy 1, Altaf A Wani 1,2
PMCID: PMC4180185  NIHMSID: NIHMS630838  PMID: 20013802

Abstract

Cisplatin is one of the most widely used anticancer agents, displaying activity against a wide variety of tumors. However, development of drug resistance presents a challenging barrier to successful cancer treatment by cisplatin. To understand the mechanism of cisplatin resistance, we investigated the role of damaged DNA binding protein complex subunit 2 (DDB2) in cisplatin-induced cytotoxicity and apoptosis. We show that DDB2 is not required for the repair of cisplatin-induced DNA damage, but can be induced by cisplatin treatment. DDB2-deficient noncancer cells exhibit enhanced resistance to cell growth inhibition and apoptosis induced by cisplatin than cells with fully restored DDB2 function. Moreover, DDB2 expression in cisplatin-resistant ovarian cancer cell line CP70 and MCP2 was lower than their cisplatin-sensitive parental A2780 cells. Overexpression of DDB2 sensitized CP70 cells to cisplatin-induced cytotoxicity and apoptosis via activation of the caspase pathway and downregulation of antiapoptotic Bcl-2 protein. Further analysis indicates that the overexpression of DDB2 in CP70 cells downregulates Bcl-2 expression through decreasing Bcl-2 mRNA level. These results suggest that ovarian cancer cells containing high level of DDB2 become susceptible to cisplatin by undergoing enhanced apoptosis.

Keywords: cisplatin, DDB2, apoptosis, Bcl-2, cisplatin resistance

The inorganic platinum (Pt) drug molecule cisplatin (cis-dia-mminedichloroplatinum [II]) was first described as an antineo-plastic agent in 1965 by Rosenberg et al.1 It has been in widespread clinical use for many years to treat various malignant tumors, including testicular, ovarian, head and neck and lung cancers.2 The major goal of cancer chemotherapy is to commit tumor cells to apoptosis after exposure to antitumor agents. Cisplatin can interact with DNA and form DNA crosslinks in target cells. These crosslinks are commonly recognized as an apoptotic stimulus.3,4 However, although cisplatin is a very potent inducer of apoptosis, resistance develops and is implied when tumor cells fail to undergo apoptosis at clinically relevant drug concentrations. This resistance can be acquired through chronic drug exposure or it can present itself as an intrinsic phenomenon.5 With the understanding that the cytotoxic effect of cisplatin is a complex process, from initial drug uptake to the final stages of apoptosis, it follows that intracellular events interfering with any stage of this process will inhibit apoptosis and lead to drug resistance.6

Cisplatin-induced apoptosis can be initiated through both intrinsic and extrinsic pathways. Cisplatin induces rapid dose-dependent release of cytochrome c from mitochondria to cyto-sol.7,8 Cytochrome c subsequently activates the caspase cascade.9 This activation leads to an irreversible commitment to apoptotic cell death. For cisplatin, caspase 3 and 9 are critical, and their activation is attenuated in resistant cells.10–13 Several members of the so-called proapoptotic (e.g., Bax, Bak, Bad and Bcl-Xs) and antiapoptotic (e.g., Bcl-2 and Bcl-XL) families of proteins reportedly regulate cisplatin-induced activation of caspase cascade. It has been observed that cisplatin-induced apoptosis in both sensitive and resistant ovarian cancer cells is associated with an increased level of Bax and Bak proteins,14 whereas downregulation of Bcl-2 or Bcl-XL sensitizes cisplatin-resistant human cancer cells to cisplatin.15–17 Cisplatin also induces apoptosis through activation of Fas/FasL system, the extrinsic apoptosis pathway.18 Therefore, cisplatin-induced apoptotic signaling likely involves several pathways, and the regulation of apoptosis signal is very complex and important in the development of cisplatin resistance.

Damage DNA binding protein complex subunit 2 (DDB2) is a 48-kDa protein originally identified as a component of the damage-specific DNA-binding heterodimeric complex DDB.19 Besides the known ability of DDB2 protein to bind ultraviolet (UV)-damaged DNA and serving as the initial damage recognition factor during nucleotide excision repair (NER),20 DDB2 was also reported to be involved in p53-mediated apoptosis on exposure to UV radiation21 and spontaneous apoptosis in testes.22 Since cisplatin-induced DNA crosslinks and UV-induced DNA damage are removed by the same NER pathway, we assessed the potential involvement of DDB2 in the repair of cisplatin-induced DNA lesions, as well as cisplatin-induced apoptosis and development of cellular resistance. Here, we show that (i) unlike the repair of UV-induced DNA damage, DDB2 is not required for the repair of cisplatin-induced DNA lesions, (ii) over expression of DDB2 can sensitize both noncancer human fibroblasts and human ovarian cancer cells to cisplatin treatment and (iii) DDB2 functions in cisplatin-induced apoptosis through downregulating the antiapoptotic protein Bcl-2 expression in human ovarian cancer cell line CP70.

Material and Methods

Cell culture and treatment

Normal human skin OSU-2 fibroblasts were established and maintained in culture as described earlier.23 Li-Fraumeni Syndrome 041 fibroblasts were originally provided by Dr. Michael Tainsky (M.D. Anderson Cancer Center, Austin, TX). This cell line harbors a codon 184 frameshift mutation in p53 gene, resulting in premature termination of translation of the p53 protein. Thus, their p53 function is null. Meanwhile, since DDB2 transcription is p53-dependent, p53 deficiency in this cell line also results in DDB2 deficiency.24 DDB2-expressing 041 cell line (041-N22) was established in our laboratory by stably transfecting pcDNA3.1-His-DDB2 into 041 cells.25 These cell lines were grown in DMEM supplemented with 10% fetal calf serum (FCS) and antibiotics (50 units/ml penicillin and 50 µg/ml streptomycin). Cisplatin sensitive human ovarian epithelial carcinoma cell line A2780 and the resistant subline CP70 were kindly provided by Dr. Paul Modrich (Duke University). Other 3 A2780-derived cisplatin-resistant cell lines MCP2, MCP3 and MCP8 were kindly provided by Dr. Tim Huang (The Ohio State University). These cell lines were cultured in RPMI 1640 medium, supplemented with 10% FCS and antibiotics. For cisplatin treatment, cells were maintained in medium with the desired doses of cisplatin (Sigma, St. Louis, MO) for 1 hr and then washed with PBS and followed by incubation in fresh drug-free medium for varying times post-treatment. For UV exposure, the cultures were washed with PBS and irradiated with UV at 10 J/m2 followed by incubation for varying times. UV-C light (254 nm) was delivered from a germicidal lamp at a dose rate of 0.5 J/m2/s, as measured by a UVX digital radiometer connected to a UVX-31 sensor (UVP, Upland, CA).

Immunoslot blotting (ISB) analysis

041 and 041-N22 cells were treated with 20 µM of cisplatin for 1 hr, washed twice with PBS and further cultured in drug-free medium for the desired time periods. The genomic DNA was isolated with phenol/chloroform/isoamyl alcohol (25:24:1) and quantified using PicoGreen kit assay (Invitrogen, Carlsbad, CA). The same amounts of denatured DNA were applied to nitrocellulose membranes and cisplatin-induced DNA crosslinks (Pt-GG) were detected with monoclonal anti-Pt-GG antibody (provided by Dr. Jűrgen Thomale, Institut fűr Zellbiologie, Universitätsklinikum Essen, Germany). The intensity of each band was quantified and the lesion concentrations were determined from a reference standards run in parallel to calculate the relative amounts of Pt-GG remaining at each time point.

Host cell reactivation assay

The host cell reactivation (HCR) assay was performed to determine the DNA repair ability of individual cell lines. For our study, the pCMV-Tag 2 expression control plasmid (containing the firefly luciferase gene, Stratagene, La Jolla, CA) was treated with cisplatin to introduce DNA damage into the plasmid DNA. Both the undamaged and the damaged pCMV-Tag 2 plasmid were then transfected into 041 and 041-N22 cells (0.5 µg/35-mm dish) using FuGene 6 transfection reagent (Roche, Indianapolis, IN). As an internal control, the pGL4.73 plasmid (Promega, Madison, WI), which carries a renilla luciferase gene, was also cotransfected into the cells. Effective repairing of cisplatin-generated DNA damage in pCMV-Tag 2 plasmid would allow expression of the luciferase gene. In contrast, if the DNA damage was not repaired because of the defects in DNA repair of the host cells, expression of the luciferase gene from the damaged pCMV-Tag 2 plasmid would be inhibited. The cells were harvested 2 days after transfection and both firefly and renilla luciferase activities were determined from the transfected cells using a Dual Luciferase Activity Detection System (Promega). The activity of firefly luciferase in each experiment was calculated as relative activity to the renilla luciferase activity to minimize the experimental variations. The ratio of luciferase activities in the same cell line for both undamaged and damaged plasmid was used to determine the DNA repair ability of the host cells.

Transfection of DDB2 cDNA

pcDNA3.1-His-DDB2 was transfected into CP70 cells using Lipofectamine 2000 (Invitrogen) according to the manufacturer’s protocol. For generation of stably transfected CP70 cells, the transfected cells were selected in the medium containing 500 µg/ml G418, and the transfectant lines stably expressing DDB2 were confirmed by Western blotting.

Growth inhibition assay

Cells were seeded in 96-well plates at an initial density of 2 × 103, incubated for 24 hr and treated with increasing doses of cisplatin for 1 hr. All experimental concentrations were replicated in quadruplicates. After the drug treatment, cultures were incubated for another 72 hr. At the end of the growth period, the cells were washed with PBS, fixed with 3.7% formaldehyde for 30 min and stained with 1.0% methylene blue for 30 min. The plate was rinsed in running water and then left to dry. One hundred microliters of solvent (10% acetic acid, 50% methanol and 40% H2O) was added to each well to dissolve the cells and optical density of the released color was read at 660 nm. The relative cell viability was calculated with the values of mock-treated cells set as 100%.

DNA fragmentation analysis

Exponentially growing cells were treated with different doses of cisplatin for 1 hr and incubated in drug-free medium for another 24 and 48 hr. At each time point, attached and detached cells were collected and DNA isolated by treatment with RNase, proteinase-K and phenol-chloroform. Cell equivalent (∼4 × 105) aliquots of purified DNA were subjected to electrophoresis on 2% agarose gels.

Flow cytometric analysis for detecting apoptotic cells

041 and 041-N22 were treated with cisplatin for 1 hr, and further cultured in drug-free medium for 48 and 72 hr. The cells were trypsinized and fixed with 70% cold ethanol. Before analyses, the fixed cells were washed with PBS and incubated with a solution containing 33 µg/ml propidium iodide, 0.2% NP-40 and 7,000 U/ml RNase A for 30 min in the dark. Flow cytometric analysis was performed using a BD FACSCalibur system equipped with a 488 nm 15 mW air cooled Argon laser (The DHLRI Flow Cytometry and Cell Sorting Analysis Core at OSU). At least 10,000 events were analyzed. Histograms were generated which indicate the relative abundance of cell populations in sub-G1, G0/G1, S and G2/M phases, and amongst them, sub-G1 represents apoptotic cells.

Colony formation assay

Exponentially growing CP70 cells were transfected with pcDNA3.1-His-DDB2 or mock-treated for 24 hr, exposed to 20 µM cisplatin for 1 hr and then reseeded in 100 mm dishes at a density of 500 cells per dish. After growing for 10 days, the cells were fixed with ethanol, stained with crystal violet and colonies of >50 cells were counted. The colony forming ability was calculated relative to the nontreated cells.

Western blotting analysis

Whole cell lysates were prepared by boiling cell pellets for 10 min in lysis buffer (2% SDS, 10% glycerol, 62 mM Tris-HCl, pH 6.8 and a complete mini-protease inhibitor cocktail [Roche Applied Science]). After protein quantification with Bio-Rad Dc Protein Assay (Bio-Rad Laboratories, Hercules, CA), equal amounts of proteins were loaded and separated on a polyacrylamide gel and transferred to a nitrocellulose membrane. Protein bands were immunodetected with appropriate antibodies, e.g., rabbit anti-DDB2 antibody generated in our laboratory,26 mouse anticleaved PARP and rabbit anti-cleaved caspase 3 antibodies from Cell Signaling Technology (Danvers, MA); mouse anti-Bcl-2, mouse anti-Tubulin, mouse anti-Actin and goat anti-Lamin B antibodies purchased from Santa Cruz Biotechnology (Santa Cruz, CA).

RT-PCR and quantitative RT-PCR

Total RNA was purified from various cell samples by using Trizol (Invitrogen). The cDNA was generated by reverse transcription using superscriptase II (Invitrogen) and oligo (dT) in a 20 µl reaction containing 1 µg of total RNA. An aliquot of 0.5 µl cDNA was used in each 20 µl PCR reaction, using PCR Master Mix (Promega). The following conditions were used: an initial denaturation at 95° C for 5 min followed by denaturation at 94° C for 30 sec, annealing at 65° C for 1 min, touchdown at − 1°C per cycle and extension at 72°C for 1 min for a total of 10 cycles. The condition was then fixed for 25 cycles of denaturation at 94° C for 30 sec, annealing at 50°C for 1 min and extension at 72°C for 1 min with a final extension at 72° C for 10 min. PCR products were analyzed by 1.5% agarose gel. For quantitative RT-PCR, we used Applied Biosystem’s Power SYBR Green PCR Master Mix and the reactions were run on an ABI 7500 Fast Real-Time PCR system. The following primers were used: DDB2, forward, 5′-CTCCTCAATGGAGGGAACAA-3′; reverse, 5′-GTG ACCACCATTCGGCTACT-3′; Bcl-2, forward, 5′-ATG TGTGTGGAGAG-CGTCAA-3′; reverse, 5′-ACAGTTCCA-CAAAGGCATCC-3′; and actin, forward, 5′-TGCCCATT-TATGAGGGCTAC-3′; reverse, 5′-GCCATCTCGTTCTCG AAGTC-3′.

Results

DDB2 expression is low in certain cisplatin-resistant cancer cells and is induced in both normal human and cancer cell lines on cisplatin treatment

To understand whether the DDB2 expression level is different in cisplatin-sensitive and -resistant cancer cells, ovarian cancer cell line A2780 (sensitive) and its derivative cisplatin-resistant cell lines CP70, MCP2, MCP3 and MCP8 were tested. All these drug-resistant cell lines were independently derived from A2780 by multiple cycles of selection with increasing cisplatin concentrations.27 As shown in Figure 1a, among four cisplatin-resistant cell lines, CP70 and MCP2 cell lines have lower DDB2 expression than A2780 cells, while the level of DDB2 in another 2 resistant cell lines, MCP3 and MCP8, is comparable to that of A2780 cells, indicating that DDB2 might be related to cisplatin resistance development in some of the cancer cell lines. Given the expression of DDB2 is regulated by p53, we wanted to know whether the DDB2 level in these ovarian cancer cell lines is correlated with p53 status. As reported previously,27 A2780 has wild-type p53 whereas CP70 has dysfunctional p53. Here, we analyzed the function of p53 in MCP2, MCP3 and MCP8 cells. As shown in Supporting Information Figure S1, both p53 and its downstream protein p21 can be induced by cisplatin in MCP3 and MPC8 cells, while not in MCP2 cells, indicating MCP3 and MCP8 have functional p53 whereas MCP2 has dysfunctional p53. Therefore, the low level of DDB2 in CP70 and MCP2 cells seems to be due to a lack of p53 function. We also evaluated the expression level of Bcl-2 in these cell lines and surprisingly found that, among the tested cell lines, those with low level of DDB2 exhibit high level of Bcl-2 and vice versa, suggesting an inverse correlation between DDB2 and Bcl-2 expression in these ovarian cancer cell lines.

Figure 1.

Figure 1

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The expression of DDB2 is low in some cisplatin-resistant ovarian cancer cells and DDB2 is induced in normal human fibroblasts and cancer cells after cisplatin treatment. (a) The whole cell lysates were prepared from cisplatin-sensitive A2780 cell line and its derivative cisplatin-resistant CP70, MCP2, MCP3 and MCP8 cell lines, the same amount of proteins were subjected to Western blotting to detect DDB2 and Bcl-2 expression, Lamin B was used as loading control. (b, c) Normal human fibroblast OSU-2 cells (b) and ovarian cancer cell line A2780 cells (c) were treated with 20 µM of cisplatin for 1 hr or UV irradiated at 10 J/m2, and further cultured in cisplatin-free medium for the indicated time periods. Cell lysates were analyzed by Western blotting with anti-DDB2 antibody. The relative amount of DDB2 at various post-treatment times were quantified relative to the respective untreated levels and normalized to Tubulin controls.

Our study and others have shown that DDB2 is degraded at early times, while induced at late times after UV irradiation.28’29 Since cisplatin-induced DNA damage is processed by the same pathway as that for the repair of UV-induced DNA damage, we wanted to know whether DDB2 follows a similar dynamics on cisplatin treatment. Normal human fibroblasts OSU-2 cells and human ovarian cancer cell line A2780 were treated with 20 µM cisplatin for 1 hr. Cells were further cultured in drug-free medium for up to 48 hr. As a control, these cells were also treated with 10 J/m2 UV, which has equivalent toxicity to 20 µM cisplatin treatment for 1 hr (data not shown), and allowed to grow for up to 48 hr. Consistent with previous studies,28 Western blotting analysis showed that UV irradiation causes DDB2 degradation at 1 and 8 hr, whereas increases the DDB2 level at 24 and 48 hr in both OSU-2 and A2780 cells (Figs. 1b and 1c, left panel). However, cisplatin treatment does not cause any obvious DDB2 degradation in either cell lines, but does show increased DDB2 level after 8-hr exposure (Figs. 1b and c, right panel). The relatively late induction of DDB2 by cisplatin may indicate its role in other cellular responses to cisplatin treatment, e.g., cisplatin-induced cellular apoptosis, but no apparent role in DNA repair

DDB2 is not required for the repair of cisplatin-induced DNA damage

Since DDB2 is not degraded after cisplatin treatment, we reasoned that DDB2 may not be involved in the repair of cisplatin-induced DNA damage. To test this hypothesis, we compared the repair efficiency of cisplatin-induced DNA damage in 041 and 041-N22 cells. As elaborated earlier, 041 cells are deficient in DDB2 expression, whereas 041-N22 cells have restored DDB2 expression by stably transfecting DDB2 in 041 cells (Fig. 2a). 041 and 041-N22 cells were treated with 20 µM of cisplatin for 1 hr, cultured further in drug-free medium for 0, 4, 8 and 24 hr. The initial and remaining cisplatin-induced DNA lesions were quantified by immunoslot blotting analysis with anti-Pt-GG antibody. As shown in Figure 2b, the removal rate of Pt-GG was comparable between 041 and 041-N22 cells, indicating that overexpression of DDB2 had no influence on the removal of cisplatin-induced DNA crosslinks. To confirm this result by another independent repair assay, the capability of 041 and 041-N22 cells to repair cisplatin-induced DNA damage was determined using HCR. Cisplatin-damaged pCMV-Tag 2 plasmid DNA was transfected into both 041 and 041-N22 cells. As a control, undamaged pCMV-Tag 2 plasmid was transfected in parallel into these cells for assessing the level of luciferase expression. HCR data show that when cisplatin-damaged pCMV-Tag 2 plasmids have been transfected into 041 or 041-N22 cells for 2 days, 85.7% and 84.5% of luciferase activity was recovered, respectively, in comparison to undamaged plasmid controls (Fig. 2c). This further confirmed that DDB2 expression is not required in the repair of cisplatin-induced DNA damage.

Figure 2.

Figure 2

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DDB2 is not required for the repair of cisplatin-induced DNA damage. (a) The expression of DDB2 in 041 and 041-N22 cells. (b) DDB2-deficient 041 and DDB2-proficient 041-N22 cells were treated with 20 µM cisplatin for 1 hr, and further cultured in drug-free medium for the indicated time periods. Total DNA was isolated and analyzed by ISB assay for cisplatin-induced DNA damage with anti-Pt-GG antibody. The relative percentage of remaining Pt-GG at different time points is an average of 3 independent repeats. Bars represent standard deviation (SD). (c) 041 and 041-N22 cells were transfected with cisplatin-damaged and undamaged pCMV-Tag 2 plasmid. As an internal control, the pGL4.73 plasmid, which carries a renilla luciferase gene, was cotransfected with the pCMV-Tag 2 plasmid. The cells were harvested 2 days after transfection and both firefly and renilla luciferase activities were determined. The activity of firefly luciferase in each experiment was calculated as relative activity to the renilla luciferase activity to minimize the experimental variations. The ratio of luciferase activities in the same cell line for both undamaged and damaged plasmid was used to determine the DNA repair ability of the host cells (n = 3, bar: SD).

DDB2 is required for cisplatin-induced cytotoxicity and apoptosis

Next, we tested the involvement of DDB2 in cisplatin-induced cytotoxicity by determining the effect of cisplatin treatment on the rate of growth inhibition in DDB2-deficient and -proficient cells. 041 and 041-N22 cells were treated with different doses of cisplatin for 1 hr and subsequently cultured for 2 days for measuring cell growth as described in “Material and Methods” section. As shown in Figure 3a, though the growth of cells was severely inhibited with increasing cisplatin concentration in both cell lines, the effect was more pronounced in DDB2-proficient 041-N22 cells than DDB2-deficient 041 cells, indicating that the presence of DDB2 exacerbates the cytotoxicity of cisplatin.

Figure 3.

Figure 3

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DDB2 is required for cisplatin-induced cellular apoptosis. (a) 041 and 041-N22 cells were treated with different doses of cisplatin for 1 hr and cells were further cultured for 48 hr. Cells were stained with methylene blue and relative cell viability was calculated. Each point represents an average of quadruplicate determinations with bars representing SD. (b, c) 041 and 041-N22 cells were treated with 80 µM cisplatin for 1 hr and further cultured in cisplatin-free medium for 48 or 72 hr. Apoptotic cells were detected using flow cytometry, and the percentage of apoptotic cells were plotted (Bar: SD, N = 3) (b). Whole cell lysates were prepared by boiling cells in SDS lysis buffer and analyzed by Western blotting (c).

Considerable evidence indicates that cisplatin kills cells through apoptotic mode of cell death.30 Therefore, we determined the effect of DDB2 on the cisplatin-induced apoptosis. 041 and 041-N22 cells were treated with 80 µM cisplatin for 1 hr and further cultured in drug-free medium for 48 and 72 hr. Apoptotic cells were first analyzed using flow cytometry. As shown in Figure 3b and Supporting Information Figure S2, the percentage of apoptotic cells induced by cisplatin treatment in 041 cells are 0.24 and 3.69% at 48 and 72 hr time points, respectively, which are elevated to 6.38 and 19.88% in 041-N22 cells. This result indicates that DDB2 is a key mediator of cisplatin-induced apoptosis. We further confirmed this finding by subjecting the whole cell lysates prepared from aforementioned cells to Western blotting. Figure 3c shows the appearance of cleaved PARP and caspase 3 in DDB2-expressing 041-N22 cells at 72 hr after cisplatin treatment, but not in DDB2-deficient 041 cells. Again, we found that DDB2 expressing 041-N22 cells have lower level of Bcl-2 than DDB2-deficient 041 cells, suggesting an inverse correlation between DDb2 and Bcl-2. These data showed an unambiguous involvement of DDB2 in invoking cisplatin-induced apoptosis. In addition, since 041 and 041-N22 are deficient in p53, it is likely that the DDB2-mediated apoptosis is independent of p53.

DDB2 is involved in overcoming cisplatin resistance

Since DDB2 participates in cisplatin-induced cytotoxicity and apoptosis in human fibroblasts, we reasoned that DDB2 might play a role in overcoming cisplatin resistance. We then investigated whether overexpression of DDB2 in CP70 cells enhances their sensitivity to cisplatin. CP70 cells were transiently transfected with DDB2 cDNA or mock-transfected for 24 hr and then treated with cisplatin for 1 hr. The clonogenic assay showed that cisplatin treatment reduced the colony formation to 46% in DDB2 overexpressing cells as compared to 98% in mock-transfected control (Fig. 4a). Cisplatin sensitivity was also measured in DDB2 stably transfected CP70 cell line. A2780, CP70, vector expressing CP70 (CP70-vector) and clone-1 of DDB2 expressing CP70 (CP70-DDB2-1) cells were treated with cisplatin and growth inhibition assessed after 3-day culture. As shown in Figure 4b, CP70-vector cells exhibited sensitivity identical to untransfected CP70 cells. Moreover, in comparison with IC50 of 6.92 µM in A2780 cells, CP70-DDB2-1 cells also exhibited enhanced cisplatin sensitivity with IC50 of 12.74 µM vs. 44.79 and 45.13 µM in untransfected CP70 and CP70-vector cells, respectively. These results indicated that DDB2 expression is instrumental in overcoming cisplatin resistance of cancer cells.

Figure 4.

Figure 4

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Overexpression of DDB2 sensitizes cisplatin-resistant CP70 cells to cisplatin treatment. (a) CP70 cells were transiently transfected with 2 µg DDB2 cDNA for 24 hr, treated with 20 µM cisplatin for 1 hr and grown in drug-free medium for 10 days. Cells were stained and colonies of >50 cells were counted. Colony formation is expressed relative to the number of colonies in control set to 1. (Bar: SD, N = 4). (b) A2780, CP70, stable vector-expressing CP70 (CP70-vector), stable DDB2-expressing CP70 (CP70-DDB2-1) cells were treated with cisplatin at varying concentrations for 1 hr and further grown in drug-free medium for 3 days. Methylene blue stained cells in the treated samples compared to untreated control cells were used to obtain relative cell survival. (Bar: SD, N = 4). (c) A2780, CP70 and CP70-DDB2-1 cells were treated with 20 µM cisplatin for 1 hr and allowed to grow for the indicated time periods. Genomic DNA was isolated and analyzed for fragmentation on 2% agarose gel.

We further tested whether overexpression of DDB2 can promote apoptosis in cisplatin-resistant CP70 cancer cells. A2780, CP70 and CP70-DDB2-1 cells were treated with 20 µM cisplatin for 1 hr and further cultured for 24 and 48 hr. The genomic DNA was isolated and resolved on aga-rose gels to detect the fragmentation of DNA, a typical hall-mark of apoptotic cell death. As shown in Figure 4c, the internucleosomal DNA fragmentation was seen at 48 hr post-treatment with cisplatin in sensitive A2780 but not in resistant CP70 cells. However, the introduction of DDB2 in CP70 cells revealed DNA fragmentation similar to that of sensitive cells. This data further corroborated the role of DDB2 in promoting cisplatin-induced apoptosis in cisplatin-resistant cancer cells.

DDB2 enhances cisplatin-induced apoptosis through downregulating Bcl-2 expression

Antiapoptotic protein Bcl-2 was reported to regulate cisplatin-induced apoptosis,10 and the Bcl-2 expression level is higher in the resistant cell line than in the sensitive cell line.31–34 More importantly, we have found that there might be an inverse correlation between DDB2 and Bcl-2 expression in human ovarian cancer cell lines (Fig. 1a). To substantiate this, we detected the Bcl-2 in CP70-DDB2-1 cells, and compared it with A2780, untransfected CP70 and CP70-vector cells at both protein and mRNA levels. Consistent with a previous report,34 Bcl-2 protein was present at a negligible level in A2780 but highly expressed in CP70 cells (Fig. 5a), which provides a basis for differential apoptosis observed in the 2 cell lines. Moreover, stable transfection of DDB2 completely depleted Bcl-2 protein in CP70 cells. The data showed a dramatic inverse correlation of DDB2 and Bcl-2 levels in these cell types. In comparison, the changes of Bcl-2 at mRNA level were consistent with the protein analysis exhibiting lower Bcl-2 mRNA in A2780 and CP70-DDB2-1 but higher Bcl-2 mRNA in CP70 and CP70-vector cells (Figs. 5b and 5c). This finding suggests that persistent DDB2 expression can down-regulate Bcl-2 by inhibiting its transcription.

Figure 5.

Figure 5

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DDB2 enhances cisplatin-induced apoptosis through downregulating Bcl-2 expression. (a) Cell lysates were prepared from A2780, CP70, CP70-vector and CP70-DDB2-1 cell lines, and analyzed by immunoblotting for the expression of DDB2 and Bcl-2. (b, c) Total RNA was isolated from cell lines described in (a) and subjected to RT-PCR and real-time RT-PCR analysis to detect the mRNA levels of DDB2 and Bcl-2. The PCR products were resolved in agarose gel (b), and the relative transcript levels obtained from quantitative RT-PCR were plotted in (c) (Bar: SD, N = 3). (d) CP70 cells were transiently transfected with 2 µg DDB2 cDNA for 24, 48 and 72 hr, whole cell lysates were prepared and analyzed by Western blotting. (e) CP70 cells were transiently transfected with 2 µg DDB2 cDNA for 24 hr, treated with 20 µM cisplatin for 1 hr and left in culture for 24 and 48 hr. Cell lysates were then prepared and analyzed by Western blotting. (f) CP70 cells were transiently transfected with 2 µg DDB2 cDNA for 24 hr, then treated with MG132 (5, 10 µM) or Bortezomib (5, 10 nM) for 24 hr. The whole cell lysates were prepared and subjected to Western blotting.

We further confirmed this finding in DDB2 transiently transfected CP70 cells. We mock or transiently transfected DDB2 cDNA into CP70 cells for 24, 48 and 72 hr. The whole cell lysates were prepared and the expression of DDB2 and Bcl-2 were analyzed using Western blotting. As shown in Figure 5d, the expression of ectopic DDB2 can sustain until 72 hr. Meanwhile, the expression of Bcl-2 reduced with the time of DDB2 transfection, further confirming overexpression of DDB2 downregulates Bcl-2 level. Furthermore, the DDB2 transiently transfected CP70 cells were treated with cisplatin for 1 hr and followed by growth in drug-free medium for 24 and 48 hr. Western blot analysis showed that cleaved PARP was detected in DDB2-expressing CP70 cells but not in untransfected CP70 cells (Fig. 5e), confirming the requirement of DDB2 in cisplatin-induced apoptosis. This result was reinforced by the concurrent detection of activated caspase 3 in CP70-DDB2 cells while very weak signals were detected in untransfected cells (Fig. 5e). Since Bcl-2 can be degraded through ubiquitin-proteasome pathway,35 we investigated the effect of DDB2 overexpression on the Bcl-2 degradation. We mock- or transiently transfected DDB2 cDNA into CP70 cells for 24 hr and then treated them with proteasome inhibitor MG132 or bortezomib or mock-treated for another 24 hr. As shown in Figure 5f, overexpression of DDB2 reduced the level of Bcl-2 (Lane 6 vs. 1), which is consistent with Figures 5d and 5e. Treatment with MG132 and bortezomib can block this reduction (Lanes 7–10 vs. 6). However, these treatments have no influence on the Bcl-2 level in CP70 cells without DDB2 overexpression (Lanes 2–5 vs. 1), indicating proteasome degradation is involved in the DDB2-mediated Bcl-2 reduction. Taken together, we concluded that DDB2 mediates cisplatin-induced apoptosis through downregulating Bcl-2 level in cells.

Discussion

DDB2 dynamics in response to cisplatin treatment

As a DNA damage recognition factor, DDB2 has been extensively studied in the process of NER after UV irradiation.36 DDB2 is recruited to UV-induced DNA damage sites; it is then ubiquitylated and degraded. The ubiquitylation and degradation of DDB2 are proposed to help targeting of another damage recognition factor XPC,37 which is indispensable in the global genomic repair of UV-induced DNA damage. Since both UV-induced DNA damage and cisplatin-induced DNA crosslinks are removed by the same DNA repair pathway, NER, DDB2 degradation would be expected to also occur in response to cisplatin treatment. However, our results do not support this scenario. The recruitment of DDB2 to the site of the DNA damage and its subsequent degradation does not seem to be universal for all types of damage. This can be explained by our finding that DDB2 is not required for the repair of cisplatin-induced DNA crosslinks. Although DDB2 has been reported to be able to bind various types of DNA damage, the binding affinity and the involvement of DDB2 in the repair of various DNA lesions are quite different (reviewed in Ref. 20). Very obvious one is the fact that UV-induced DNA damage pyrimidine (6-4) pyrimidine photoproducts (6-4PP) has high affinity to DDB2 in vitro, but its repair does not require DDB2; while the repair of cyclobutane pyrimidine dimers (CPD), which induces relatively minor DNA structural distortion, does require DDB2.20 Since cisplatin-induced DNA crosslinks cause a major DNA helix distortion compared to CPD or even 6-4PP,38–40 it is understandable that the repair of cisplatin-DNA crosslinks does not require DDB2 binding and consequently DDB2 degradation would not be expected either.

It has been reported that DDB2 is induced after 24–48 hr post-UV irradiation, and our result also confirmed this finding. However, cisplatin-dependent DDB2 induction occurs as early as 8 hr post-treatment, suggesting that DDB2 induction is more important than DDB2 degradation in the cellular events after cisplatin treatment. Among the DNA damage responses induced by cisplatin treatment, apoptosis is a relatively late event. Given our finding that DDB2 is required for cisplatin-induced apoptosis, we reckon that the induction of DDB2 is for regulating apoptosis after cisplatin treatment.

DDB2 regulates cisplatin-induced apoptosis independently of p53

p53 is clearly a major player in the apoptotic response of cells. Many tumor cell lines that have been studied to date undergo apoptosis following DNA damage induction to a greater extent if they express wild-type p53. In addition, DDB2 and p53 are also shown to crossregulate each other.41 Human DDB2 expression is positively regulated by p53 at both the basal level and after DNA damage.24 As a downstream effecter of p53, DDB2 recognizes UV-induced DNA damage and stimulates NER.42 On the other hand, DDB2 was found to directly regulate p53 level and its downstream-regulated proteins before and after UV irradiation. Importantly, apoptotic cell killing and caspase 3 induction by UV irradiation were clearly restored to the DDB2-deficient XP-E strain by the p53 cDNA expression41 indicating that DDB2 can participate in UV-induced apoptosis through regulating p53. However, we found that DDB2 can also enhance cisplatin-induced apoptosis in the absence of p53. 041 fibroblasts have mutated p53,43 and CP70 cells were reported to possess a dysfunctional p53.27 In our studies, both cell lines exhibited enhanced cisplatin-induced apoptosis on DDB2 overexpression, indicating that the DDB2 involvement in apoptosis is direct and independent of cellular p53 status.

Our results indicated that DDB2 can downregulate the level of antiapoptotic protein Bcl-2 at both mRNA and protein level. It has been reported that the transcription of Bcl-2 is E2F1-dependent.44 Given that DDB2 can interact with transcription factor E2F1,45 we then reason that DDB2 may influence Bcl-2 transcription through interaction with E2F1 and further downregulating its transcription activity. At post-translational level, Bcl-2 can be modified by ubiquitylation and then degraded by proteasome.35 Since DDB2 is a subunit of DDB-Cul4A E3 ubiquitin ligase complex,46 we speculate that DDB2 may also downregulate Bcl-2 through promoting its ubiquitylation and degradation. However, these views require future experimental proof.

DDB2 deficiency offers one of the mechanisms for acquired resistance of ovarian cancers to cisplatin treatment

We have tested the DDB2 expression in several cisplatin-resistant human ovarian cancer cell lines and found 2 of them have low level of DDB2 compared to their parent cisplatin-sensitive cell line. On the basis of our strong data that the protein level of DDB2 inversely correlates with Bcl-2, we believe that DDB2 deficiency is instrumental in the development of cisplatin resistance in human ovarian cancer cells. However, with the understanding that the cytotoxic effect of cisplatin is a complex process, from initial drug uptake to the final stages of apoptosis, it is logical that intracellular events interfering with any stage of this process will inhibit apoptosis and lead to drug resistance. Thus, DDB2 deficiency is not the single determinant of cisplatin resistance and high DDB2 expression does not always indicate cisplatin sensitivity, as evidenced by the finding that the DDB2 expression is not low in cisplatin resistant MCP3 and MCP8 cell lines. Interestingly, in our DDB2 stably transfected CP70 cells, we surprisingly found that besides the exogenous DDB2, the endogenous DDB2 level was also increased. Therefore, we speculate that the transfected exogenous DDB2 competes with the endogenous DDB2 for degradation to cause the accumulation of endogenous DDB2. Furthermore, this finding indicates that the low-protein level of DDB2 in cisplatin-resistant CP70 cells can be induced in the absence of functional p53. Given the fact that p53 is frequently mutated in tumors, the p53-independent DDB2 induction in cisplatin-resistant cancer cells can provide a novel strategy to enhance cisplatin-induced apoptosis and to overcome cisplatin resistance in p53 mutated tumors. In conclusion, our studies have linked DDB2 to cisplatin sensitivity and Bcl-2 expression. On the basis of our data, we posit that cisplatin treatment induces DDB2 expression. Subsequently, increased DDB2 inhibits the transcription of antiapoptotic protein Bcl-2. Meanwhile, DDB2 enhances the proteasome-mediated Bcl-2 protein degradation. As a result, cellular Bcl-2 level decreases and cisplatin damaged DNA-mediated apoptosis is enhanced (Fig. 6). Cisplatin resistance is known to develop through alterations of several different cellular pathways, and our study presents a very significant addition to this repertoire. Since our study is based on a single noncancer and a single cancer parental cell line, more cell lines will have to be investigated to reaffirm the universality and implications of this unique finding. It is evident, however, that just as all the known cisplatin resistance mechanisms cannot exist in a single cancer type, the DDB2 deficiency-based cisplatin resistance cannot be operational in all the cisplatin resistant cancers.

Figure 6.

Figure 6

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Schematic presentation of the role of DDB2 in cisplatin-induced apoptosis. DDB2 is induced by cisplatin treatment. Subsequently, increased DDB2 inhibits Bcl-2 transcription and enhances Bcl-2 protein degradation, causing the downregulation of cellular Bcl-2 level. As a result, cisplatin-induced apoptosis is promoted. [Color figure can be viewed in the online issue, which is available at www.interscience.wiley.com.]

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Acknowledgements

This work was supported by NIH grants ES2388, ES12991 (A.A.W.) and CA93413 (A.A.W. and Q.-E.W.). B.B., a visiting scholar from Al-Azhar University, was supported by an Egyptian Channel Fellowship. The authors thank Drs. Michael Tainsky, Paul Modrich and Tim Huang for providing cell lines, Dr. Jűrgen Thomale for anti-Pt-GG antibody and Dr. Yi-Wen Huang for help with real-time PCR analysis.

Footnotes

Additional Supporting Information may be found in the online version of this article.

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