Hypoxia Promotes Synergy between Mitomycin C and Bortezomib through a Coordinated Process of Bcl-xL Phosphorylation and Mitochondrial Translocation of p53 Free

Peritoneal carcinomatosis is regarded as the most common lethal primary or secondary cancerous disease that affects the peritoneal cavity following colorectal cancer, appendiceal cancer, ovarian cancer, gastric cancer, or diffuse malignant peritoneal mesothelioma (1, 2). Colorectal peritoneal carcinomatosis (CPC), which affects approximately 10% of colorectal cancer patients, is one manifestation of metastatic colorectal cancer (2, 3). Standard treatments involve cytoreductive surgery and perioperative chemotherapy (4). However, CPC is generally considered an untreatable condition that makes clinicians abandon further aggressive treatment, and the response rates of CPC patients are low (4). Physiologic stress from hypoxia, a feature of most tumors, has been associated with the resistance of CPC to therapy (5).

Bioreductive prodrugs have been designed to be activated under hypoxia and thus increase achievable therapeutic effects under hypoxic conditions. Some agents have also been developed to target hypoxic tumor cells through inhibition of hypoxia-inducible factor (HIF) 1, a master transcriptional regulator (6). The chemotherapeutic agent mitomycin C is a naturally occurring, clinically used, bioreductive alkylating prodrug that is activated by several enzymes, including cytochrome P450 reductase (7). Mitomycin C has also been shown to inhibit HIF-1α protein synthesis (8). It has been tested in a great number of preclinical and clinical studies and is widely used to treat peritoneal carcinomatosis (9). However, mitomycin C alone is not potent enough to generate sustained tumor regression and has been shown cause unexpected side effects (10, 11). Therefore, we should consider combinations of cytotoxic drugs and other agents that have achieved improvement in animal models and patients (12–14), thereby enhancing the likelihood of survival for CPC patients.

The ubiquitin–proteasome pathway has been well recognized as a valid target for antitumor therapy and is also regarded as a biomarker of poor prognosis for peritoneal mesothelioma tumors (15). Bortezomib, an approved-for-clinical-use proteasome inhibitor used to treat multiple myeloma, induces apoptosis in a wide variety of cancer cell lines, yet has relatively few toxic effects on normal cells (16–18). However, bortezomib has minimal activity as a single agent in the treatment of recurrent platinum-sensitive epithelial ovarian or primary peritoneal cancer, indicating that proteasome inhibitors should be combined with other reagents to increase the effectiveness of therapy (19, 20).

In this study, we observed that the combination of the chemotherapeutic agent mitomycin C and the proteasome inhibitor bortezomib in sublethal doses induced synergistic cytotoxicity and apoptosis, in particular under hypoxia in vitro and in vivo. The combination of mitomycin C and bortezomib induced oligomerization of Bcl-2 homologous antagonist/killer (Bak) through coordinated action of Bcl-xL dissociation from Bak and p53 mitochondrial localization and resulted in mitochondria-dependent apoptotic cell death. The significance of our findings promotes the opportunity to incorporate bortezomib with mitomycin C to preferentially target not only normoxic, but also hypoxic tumor cells, allowing for more effective therapy for CPC patients.

Human colorectal carcinoma CX-1 cells, which were obtained from Dr. J.M. Jessup (Division of Cancer Treatment and Diagnosis, NCI, NIH, Bethesda, MD), were cultured in RPMI1640 medium containing 10% FBS. Human colorectal carcinoma HCT116 cells, kindly provided by Dr. B. Vogelstein (Johns Hopkins University, Baltimore, MD), were cultured in McCoy's 5A medium containing 10% FBS. LS174T and LS180 are human colon adenocarcinoma cell lines and were purchased from ATCC and cultured in Eagle's minimum essential medium and 10% FBS at 37°C and 5% CO₂. Mycoplasma testing was conducted routinely for all cell lines. For transient transfection, cells were transfected with Lipofectamine 2000 (Life Technologies) and were treated 48 hours after transfection.

Hypoxic cell culture experiments were carried out at 1% O₂ using a hypoxia chamber (Anaerobic System model 1025/1029, ThermoForma).

Mitomycin C was obtained from Santa Cruz Biotechnology. Bortezomib was obtained from Millennium Pharmaceuticals. Anti-hemagglutinin (HA), anti-Bcl-xL, anti-carcinoembryonic antigen (CEA), anti-phospho-c-Jun NH₂-terminal kinase (JNK), anti-JNK, anti-Bak, anti-caspase-8, anti-caspase-9, anti-caspase-3, anti-Bad, anti-p-p53(Ser46)/p53, anti-COX IV antibody, and anti-PARP antibody came from Cell Signaling Technology. Anti-actin antibody came from Sigma-Aldrich. Anti-p-Bcl-xL (S62) antibody was obtained from Millipore. Anti-cytochrome c antibody came from BD PharMingen. Anti-Ki67 antibody was purchased from Dako.

MTS studies were carried out using the Promega CellTiter 96 AQueous One Solution Cell Proliferation Assay (Promega). Cells were then treated with MTS/phenazine methosulfate solution for 1 hour at 37°C. Absorbance at 490 nm was determined using an ELISA plate reader.

Cells were harvested with trypsinization, washed with serum-free medium, and suspended in binding buffer [annexin V-fluorescein isothiocyanate (FITC) Staining Kit, BD PharMingen]. This cell suspension was stained with FITC-conjugated annexin V and propidium iodide (PI) and then immediately analyzed using flow cytometry.

Combination index (CI) was calculated using the CompuSyn software (ComboSyn, Inc.). On the basis of CI values, the extent of synergism/antagonism was determined. In general, CI values below 1 suggest synergy, whereas CI values above 1 indicate antagonism between the drugs. CI values in the 0.9 to 1.10 range mainly indicate additive effects, those between 0.9 and 0.85 suggest slight synergy, those in the range of 0.7 and 0.3 indicate moderate synergy, and those less than 0.3 suggest strong synergy.

Cells were lysed with Laemmli lysis buffer and boiled for 10 minutes. Protein content was measured with BCA Protein Assay Reagent (Thermo Scientific), separated by SDS-PAGE, and electrophoretically transferred to nitrocellulose membrane. The nitrocellulose membrane was blocked with 5% nonfat dry milk in PBS-Tween-20 (0.1%, v/v) for 1 hour and incubated with primary antibody at room temperature for 2 hours. Horseradish peroxidase–conjugated anti-rabbit or anti-mouse IgG was used as the secondary antibody. Immunoreactive protein was visualized using the chemiluminescence protocol. Gel densitometry was analyzed using the software Gel-pro application from Media Cybernetics.

Briefly, cells were lysed in CHAPS lysis buffer with protease inhibitor cocktail (Calbiochem). A total of 0.5–1 mg of lysate was incubated with 1.5 μg of anti-Bak/HA antibody or rabbit IgG (Santa Cruz Biotechnology) at 4°C overnight, followed by the addition of protein G PLUS-agarose beads (Santa Cruz Biotechnology) and rotation at room temperature for 2 hours followed by immunoblot analysis.

Cells were pelleted and resuspended in Hoechst buffer. The cell suspension was homogenized, and spun at 1,000 × g for 15 minutes at 4°C. The supernatant was transferred and spun at 10,000 × g for 15 minutes at 4°C to pellet mitochondria. Aliquots of isolated mitochondrial fractions and cytosolic fractions were cross-linked with 1 mmol/L dithiobis (Pierce). Samples were subjected to SDS-PAGE under nondenaturing conditions followed by immunoblotting for Bak.

Cells were stained with 300 nmol/L MitoTracker (Life Technologies). Cells were washed three times with 0.5% bovine serum albumin in PBS, followed by fixation in 4% paraformaldehyde for 15 minutes. Bak was stained with anti-Bak antibody. Nuclei were stained with 4′,6-diamidino-2-phenylindole (DAPI; Cell Signaling Technology). Slides were mounted and visualized in 0.4 μm sections using an OlympusFluoview 1000 confocal microscope and the companion software FV10-ASW2.1 under a 63X oil immersion objective.

To determine the release of cytochrome c from the mitochondria, mitochondrial and cytosolic fractions were prepared using the Mitochondrial Fractionation Kit (Active Motif) following company instructions and reagents included in the kit. Cytosolic fractions were subjected to SDS-PAGE and analyzed with immunoblotting using anti-cytochrome c antibody.

Cells were transfected with 50 nmol/L of p53 siRNA and control siRNA from Sigma Aldrich, using Lipofectamine 2000. Target sequences for preparing siRNAs of human p53 were as follows: 5′-GAC UCC AGU GGU AAU CUA C-3′ (sense strand) and 5′-GUA GAU UAC CAC UGG AGU C-3′ (complement strand). Expression levels were determined by immunoblot analysis.

Nude mice (6-week-old, Taconic) were intraperitoneally (i.p.) challenged with 5 × 10⁵ LS174T cells expressing firefly luciferase, which was stably transduced by lentiviral transfection of the pGL4 Luciferase Reporter Vector (Promega) and selected with puromycin. The luciferase signal was monitored by injecting the luciferase substrate luciferin (150 mg/kg, i.p.; GoldBio) 5 minutes after anesthesia with 2% isoflurane prior to imaging on an IVIS200 system (Perkin Elmer). Bioluminescence signal was quantified using the LivingImage software (Perkin Elmer). The mice were examined for bioluminescent signals, indicating basal tumor loading. Mice with the same bioluminescence levels were divided into four groups (five per group) and subjected to different treatments, including control, bortezomib (0.5 mg/kg, i.p.) alone, mitomycin C (1.5 mg/kg, i.p.) alone, or bortezomib and mitomycin C in combination. Each group of mice was treated every other day. The tumor load, represented by bioluminescent signal determined using the IVIS Bioluminescent Imaging System, was checked from week 1 to week 5. Mice were fed ad libitum and maintained in environments with controlled temperatures of 22°C to 24°C and 12-hour light and dark cycles. All animal experiments were carried out at the University of Pittsburgh in accordance with the Guide for the Care and Use of Laboratory Animals.

For immunohistochemical staining, tumor tissues recovered from each group were fixed in 10% neutral buffered formalin and embedded in paraffin. Sections were cut 4 μm thick and then were stained either via standard hematoxylin and eosin (H&E) staining or immunohistochemical staining with mAbs specific for human CEA and Ki67 and were visualized with the Envision + dual link system (Dako) following the manufacturer's instructions.

Pimonidazole (60 mg/kg, Hypoxyprobe, Inc.) was administered 60 minutes intraperitoneally prior to sacrifice of the mice. For quantification of hypoxic tumor cell apoptosis via immunofluorescence, tumors were fixed in 4% paraformaldehyde, 30% sucrose overnight, and then frozen in optimum cutting temperature (OCT) and cryosectioned. Sections were incubated with anti-cleaved caspase-3 antibody and mouse anti-pimonidazole Dylight549-Mab, and then incubated with anti-Rabbit Alexa Fluor 488. The percentage of cleaved caspase-3–positive cells was quantified by counting at least 300 cells for each sample and plotted as mean ± SD of two independent experiments.

Pictures were taken using an Olympus BH2 microscope (Olympus) with a camera (Coolsnap3.3; Photometrics). Tissue sections of inoculation sites were examined under an Olympus BX71 microscope (Olympus) equipped with Hoffman objective lenses (×40 or 20; Diagnostic Instruments) connected to a SPOT One Digital Camera (Diagnostic Instruments). Images were then processed using Adobe Photoshop software (Adobe).

Statistical analysis was carried out using GraphPad InStat 3 software (GraphPad Software, Inc.). Data showing comparisons between two groups were assessed using the Student's t test. Comparisons among more than two groups were done using ANOVA with the appropriate post hoc testing. Statistical significance is marked with asterisks (*, P < 0.05; ** or ##, P < 0.01).

First, we investigated the effect of mitomycin C and bortezomib at different doses on human colorectal HCT116, CX-1, and LS174T cancer cells. We observed that the hallmark feature of apoptosis, PARP cleavage, significantly increased with the combination treatment of mitomycin C and bortezomib in a dose-dependent manner in all tested cell lines (Fig. 1A). We then tested the cytotoxicity of the combination of mitomycin C and bortezomib in LS174T cells under both normoxic (21% O₂) and hypoxic (1% O₂) conditions using MTS assay. As shown in Fig. 1B, mitomycin C induced 17.1% ± 2.1% cell death under normoxia. Interestingly, mitomycin C induced 29.1% ± 3.4% cell death under hypoxia (P < 0.05). Bortezomib at 10 nmol/L barely induced cell death under normoxia as well as hypoxia. Notably, bortezomib enhanced the effect of mitomycin C under both normoxia and hypoxia. To clarify whether the cytotoxicity of the combination of mitomycin C and bortezomib was associated with apoptosis, flow cytometry assay Annexin/PI was performed. We observed that the combination of mitomycin C and bortezomib induced greater apoptotic cell death (annexin V⁺ PI⁻ + annexin V⁺ PI⁺) than any single drug treatment under both normoxia and hypoxia (Fig. 1C). We also analyzed the synergistic effect of the combination of mitomycin C and bortezomib under hypoxic conditions using CI as shown in Table 1. CI values were less than 0.7 or 0.3 for the indicated doses of each drug under hypoxia, indicating moderate synergy or strong synergy.

Next, we examined whether the combination of mitomycin C and bortezomib promoted the caspase pathways under normoxia and hypoxia in human colorectal LS174T and LS180 cancer cells. It is well known that the intracellular level of hypoxia-inducible factor-1α (HIF-1α) is elevated under hypoxia due to inhibition of degradation (21–23). Indeed, data from immunoblot analysis demonstrated an elevation of HIF1α under hypoxia in LS174T cells (inset of Fig. 2A). We also observed that 2 μg/mL mitomycin C or 10 nmol/L bortezomib alone had no significant apoptotic effect under normoxia (main of Fig. 2A), but the combination of mitomycin C and bortezomib resulted in significant activation of caspase-8, 9, and 3, as well as PARP cleavage. Interestingly, mitomycin C alone under hypoxia-induced activation of caspases and PARP cleavage, while the activation and cleavage from the combination of mitomycin C and bortezomib was greatly increased. Similar results were obtained in LS180 cells (Fig. 2C).

To confirm the differential effect of mitomycin C under normoxia and hypoxia, LS174T cells were treated with various doses of mitomycin C. As shown in Fig. 2B, the effect of mitomycin C was significantly increased under hypoxia compared with that under normoxia, especially in low doses.

We next investigated the apoptotic death-related MAPK signaling pathways during treatment with combination of mitomycin C and bortezomib under normoxia and hypoxia (Figs. 3A and B). We observed phosphorylation (activation) of JNK as well as p38 along with Bcl-xL phosphorylation during treatment with the combination of mitomycin C and bortezomib, especially under hypoxic conditions in LS174T and LS180 cells. We also observed p53 elevation with mitomycin C treatment regardless of oxygen tensions in LS174T and LS180 cells. It is reported that activated p38 can phosphorylate p53 at Ser46 and phosphorylated p53 (p-p53) localizes to the mitochondria (24–27). Thus, we examined p-p53 (Ser46) and observed an increase in the level of p-p53 during treatment with mitomycin C, in particular under hypoxia, in both LS174T and LS180 cells (Figs. 3A and B).

Several researchers have reported that MAPK plays a critical role in the death receptor–initiated extrinsic as well as the mitochondrial intrinsic apoptotic pathways (28, 29). Pretreatment of SB203580, a p38 inhibitor, or SP600125, a JNK inhibitor, attenuated the drug combination–induced PARP cleavage and decreased Bcl-xL phosphorylation in both normoxic and hypoxic culture systems (Fig. 4A). These results suggest that activation of p38 and JNK plays a role in the drug combination treatment–induced apoptosis through phosphorylation of Bcl-xL. Our previous studies demonstrated that stress-induced Bcl-xL phosphorylation occurs on serine 62 (S62) residue (30). We next examined the role of Bcl-xL phosphorylation at S62 in the drug combination treatment–induced apoptosis. LS174T cells were transiently transfected with a sham plasmid, wild-type Bcl-xL (Bcl-xL WT-HA) or phospho-defective Bcl-xL (Bcl-xL S62A-HA) and then treated with mitomycin C and bortezomib under normoxia and hypoxia. We observed that Bcl-xL-S62A reduced the drug combination–induced PARP cleavage compared with Bcl-xL-WT (Fig. 4B, left and right). These data suggest Bcl-xL phosphorylation at S62 is essential for the synergistic effect of mitomycin C- and bortezomib-induced apoptosis.

To further study the role of Bcl-xL phosphorylation at S62 in the synergistic effect of mitomycin C- and bortezomib-induced apoptosis, we examined the interaction between Bak and Bcl-xL during treatment with mitomycin C and bortezomib. Bcl-xL-WT and Bcl-xL-S62A transient transfected cells were employed and immunoprecipitation assay was conducted. Figure 4C shows that Bcl-xL WT dissociated from Bak during combined treatment with mitomycin C and bortezomib under both normoxia and hypoxia, whereas Bcl-xL-S62A still maintained the interaction with Bak during combined treatment with mitomycin C and bortezomib under both normoxia and hypoxia. These results suggest that phosphorylation of Bcl-xL at S62 leads to apoptosis through dissociation of Bcl-xL from Bak.

We then further investigated the role of p53 in apoptotic death during treatment with mitomycin C and bortezomib under normoxia and hypoxia. As shown in Fig. 5A, we observed that p53 knockdown abrogated the combinatorial treatment-induced PARP cleavage under both normoxia and hypoxia, suggesting the proapoptotic function of p53. In Fig. 5B, we also observed that p53 translocated to the mitochondria during treatment with mitomycin C under normoxia, and this effect was enhanced under hypoxia probably due to p53 phosphorylation at Ser46 (Figs. 3A and B). There is no difference in terms of p53 translocation during bortezomib treatment under normoxia compared with that under hypoxia. Consistently, cellular cytochrome c release from the mitochondria to the cytosol increased in the combinatorial treatment under hypoxia compared to that under normoxia. Data from confocal immunofluorescence microscopy assay (Fig. 5C) confirmed p53 accumulation in the nucleus and mitochondria during mitomycin C alone and the combinatorial treatment, in particular under hypoxia.

Next, we investigated the role of mitochondrial translocated p53. As shown in Fig. 5D, in the LS174T cell line, p53 bound to Bak during the combinatorial treatment of mitomycin C and bortezomib. Binding affinity increased during treatment with mitomycin C alone or the combinatorial treatment under hypoxia. We also observed the dissociation of Bcl-xL from Bak during the bortezomib treatment as well as during the combinatorial treatment, especially under hypoxia, which was due to Bcl-xL phosphorylation. Similar results were obtained in the LS180 cell line. The dissociated (activated) proapoptotic members Bak or Bcl-2–associated X protein (Bax) are well known to form pores in the mitochondrial outer membrane, which allows cytochrome c to release into the cytoplasm and induce caspase cascade (31). LS174T, a Bax-null cell line (32), is a good model to study the role of Bak. Consistently, we observed increased multimeric Bak oligomerization during the treatment with mitomycin C and bortezomib, especially under hypoxia (Fig. 5E). Confocal assay also shows Bak clustering localized in the mitochondria during the combinatorial treatment, especially under hypoxia (Fig. 5F).

Finally, in vivo studies were performed to examine the effect of the combinatorial treatment with mitomycin C and bortezomib on the growth of LS174T intraperitoneal tumors. Figure 6A and B show that administration with bortezomib every other day at doses of 0.5 mg/kg significantly decreased tumor growth compared with the control group (P < 0.01). Mitomycin C alone caused a statistically significant decrease of tumor growth (P < 0.01). In particular, the combinatorial treatment with mitomycin C and bortezomib was significantly more effective in inhibiting tumor growth compared with single treatment (P < 0.01). In Fig. 6C, H&E and CEA (used as a tumor marker) staining show that all the sections were in the tumor tissue. We also observed a decrease in Ki67 expression during treatment with mitomycin C and bortezomib, especially in the combinatorial treatment (Fig. 6C). In Fig. 6D and E, tumor tissue was stained for detection of both cleaved caspase 3 (green) and pimonidazole adducts (red), and analyzed (inset of Fig. 6D) and then histograms were drawn. We observed increased cleaved caspase-3 staining in the combinatorial treatment compared with single treatment (P < 0.05). Notably, we also observed that mitomycin C and the combinatorial treatment had more cleaved caspase-3 staining under hypoxia (P < 0.05). Figure 6F shows a schematic diagram that explains how the combinatorial treatment with mitomycin C and bortezomib effectively induced apoptosis under hypoxia through Bak activation (oligomerization), Bcl-xL dissociation (Bcl-xL phosphorylation) and p53 translocation to the mitochondria.

Malignant tumors contain hypoxic regions due to insufficient blood supply and can adapt and survive under hypoxic conditions with increased survival signaling and loss of apoptotic potential (33). Given the central role of hypoxia in tumor progression and resistance to therapy, it is worth exploiting validated treatments for targeting hypoxia for therapy of CPC. In this study, we observed that a combinatorial treatment with the bioreductive prodrug mitomycin C and the proteasome inhibitor bortezomib synergistically induced apoptosis in colorectal cancer cells and the peritoneal animal model under both normoxic and hypoxic conditions, and we also investigated the underlying mechanisms of mitomycin C- and bortezomib-induced apoptosis under hypoxia.

There are two main approaches for targeting hypoxia: bioreductive prodrugs and inhibitors of molecular targets upon which hypoxic cell survival depends. Prodrugs include five different chemical moieties (nitro groups, quinones, aromatic N-oxides, aliphatic N-oxides, and transition metals), which have the potential to be metabolized by enzymatic reduction under hypoxic conditions (7). The quinone mitomycin C is preferentially activated in hypoxic tumor cells (34). However, the detailed mechanisms of how mitomycin C selectively induces the apoptotic death of hypoxic colorectal cancer cells have been poorly studied. In this study, we observed that p53 preferentially translocated to the mitochondria under hypoxia during treatment with mitomycin C (Fig. 5C).

The tumor suppressor gene p53 has multiple functions, including cell-cycle regulation, DNA repair, and apoptosis. p53-associated apoptotic regulation can be achieved in a transcription-dependent or -independent manner. Death signals have been reported to be able to induce p53 translocation to the mitochondria via the transcription-independent apoptotic signal (35–38). p53 participates directly in the intrinsic apoptosis pathway by interacting with the multidomain members of the Bcl-2 family to induce mitochondrial outer membrane permeabilization, regardless of several common mutations (39–41). In the current studies, we observed that for mitomycin C, the combinatorial treatment, as well as hypoxic conditions, induced translocation of p53 protein to the mitochondria, while the application of bortezomib had a limited effect on p53 elevation and mitochondrial translocation. Recently, some novel molecules, such as Tid1 (42), anionic phospholipids (36), dynamin-related protein 1 (43), DJ-1 (44), etc., have been reported to regulate the function of p53 translocation to the mitochondria. The possible involvement of these molecules in the mitochondrial translocation of p53 during treatment with mitomycin C and bortezomib, especially under hypoxia, needs to be elucidated in the near future.

It is obvious that mitomycin C alone is not enough to trigger sustained apoptosis, even with the dramatic translocation of p53 to the mitochondria upon hypoxia, which strongly indicates that a combination of other drugs is needed to induce conformational activation of pro-apoptotic Bak. We reported here that bortezomib enhanced the apoptotic effect of mitomycin C in several different colorectal cancer cells under both normoxic and hypoxic conditions. We propose that further liberation of Bak, specifically from the p53-activated Bak–Bcl-xL complex, is a prerequisite for inducing robust apoptosis.

We previously reported that Bcl-xL, a key antiapoptotic molecule, undergoes phosphorylation in several different treatments (30, 45, 46). We report here that activation of p38 and JNK are both specifically required for the combinatorial treatment with mitomycin C and bortezomib, which is in agreement with other observations (28, 47). The fact that a validated phospho-defective Bcl-xL mutant was resistant to the combinatorial treatment of oxaliplatin and bortezomib-induced apoptosis and retained the interaction between Bak and Bcl-xL, indicated that Bcl-xL phosphorylation at Ser 62 is a key regulatory mechanism for antagonizing antiapoptotic function in the combinatorial treatment. Thus, the mode of action of the synergistic effect of the DNA-damaging drug mitomycin C and proteasome inhibitor bortezomib is mediated via p53 mitochondrial translocation and phosphorylation of Bcl-xL which results in facilitating the disruption of the mitochondrial apoptosis effector Bak. Our results support a concept that coordinated cell-intrinsic death signals enhances in robust apoptosis.

Taken together, we presented here a novel combinatorial treatment with the chemotherapeutic agent mitomycin C and the proteasome inhibitor bortezomib, which induced synergistic apoptosis under both normoxic and hypoxic conditions through p53 translocation to the mitochondria and Bcl-xL phosphorylation, and further disruption of the mitochondrial apoptosis effector Bak. Mitomycin C and bortezomib are both commonly used drugs that are approved by the FDA and could be considered for CPC patients in clinics in the near future.

No potential conflicts of interest were disclosed.

Conception and design: X. Song, Y.T. Kwon, Y.J. Lee

Development of methodology: Y.T. Kwon, Y.J. Lee

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): X. Song, A.-K. Dilly, H.A. Choudry, Y.J. Lee

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): X. Song, H.A. Choudry, Y.J. Lee

Writing, review, and/or revision of the manuscript: X. Song, D.L. Bartlett, Y.J. Lee

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): Y.T. Kwon, Y.J. Lee

Study supervision: H.A. Choudry, Y.J. Lee

The authors thank Dr. Timothy C. Chambers (University of Arkansas for Medical Sciences, Little Rock, AR) who provided pBcl-xL-WT and pBcl-xL-S62A.

This work was supported by the following grants: NCI grant R01CA140554 (to Y.J. Lee) and Basic Science Research Program of the National Research Foundation of Korea funded by the Ministry of Science, ICT and Future Planning: NRF-2013R1A2A2A01014170 (to Y.T. Kwon). This project used the UPCI Core Facility and was supported in part by award P30CA047904.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.