Combination chemotherapy is standard for metastatic colorectal cancer; however, nearly all patients develop drug resistance. Understanding the mechanisms that lead to resistance to individual chemotherapeutic agents may enable identification of novel targets and more effective therapy. Irinotecan is commonly used in first- and second-line therapy for patients with metastatic colorectal cancer, with the active metabolite being SN38. Emerging evidence suggests that altered metabolism in cancer cells is fundamentally involved in the development of drug resistance. Using Oncomine and unbiased proteomic profiling, we found that ATP citrate lyase (ACLy), the first-step rate-limiting enzyme for de novo lipogenesis, was upregulated in colorectal cancer compared with its levels in normal mucosa and in chemoresistant colorectal cancer cells compared with isogenic chemo-naïve colorectal cancer cells. Overexpression of exogenous ACLy by lentivirus transduction in chemo-naïve colorectal cancer cells led to significant chemoresistance to SN38 but not to 5-fluorouracil or oxaliplatin. Knockdown of ACLy by siRNA or inhibition of its activity by a small-molecule inhibitor sensitized chemo-naïve colorectal cancer cells to SN38. Furthermore, ACLy was significantly increased in cancer cells that had acquired resistance to SN38. In contrast to chemo-naïve cells, targeting ACLy alone was not effective in resensitizing resistant cells to SN38, due to a compensatory activation of the AKT pathway triggered by ACLy suppression. Combined inhibition of AKT signaling and ACLy successfully resensitized SN38-resistant cells to SN38. We conclude that targeting ACLy may improve the therapeutic effects of irinotecan and that simultaneous targeting of ACLy and AKT may be warranted to overcome SN38 resistance. Mol Cancer Ther; 12(12); 2782–91. ©2013 AACR.
Current standard combination chemotherapy regimens for patients with metastatic colorectal cancer contain 5-fluorouracil (5-FU) in combination with oxaliplatin or irinotecan. Although the response rate to systemic therapies is about 50%, drug resistance develops in nearly all patients with metastatic colorectal cancer, leading to about 50,000 deaths each year in the United States (1). Advances in targeted therapy have not dramatically improved the outcomes of patients with metastatic colorectal cancer, and the majority of those with unresectable metastatic disease still die within 2 years of the diagnosis of metastasis (2). A better understanding of the mechanisms by which cancer cells develop resistance to individual chemotherapeutic agents is urgently needed to identify novel therapeutic targets and methods that would improve survival.
The chemotherapeutic agent irinotecan [CPT-11 (Camptosar)], which is commonly used in treating cancers such as metastatic colorectal cancer, causes S-phase–specific death of proliferating cells by inhibiting DNA topoisomerase I (3, 4). Irinotecan is converted by carboxylesterase into SN38 (Supplementary Fig. S1), which is 1,000-fold more potent than unconverted irinotecan (5). Cancer cells may resist SN38-mediated cell death by various mechanisms: upregulation of drug efflux (6, 7), reduction of topoisomerase I levels (8, 9), development of mutations in topoisomerase I (10), enhancement of DNA repair (11), and activation of NF-κB–regulated pathways (12). Emerging evidence suggests that altered metabolism in cancer cells is fundamentally involved in the development of drug resistance (13–15). Many of the above mechanisms may result from a reprogramming of cellular metabolism, one of the hallmarks of cancer, as in enhanced aerobic glycolysis, glutaminolysis, and de novo lipogenesis (16–18). The targeting of key metabolic enzymes sustaining these cancerous metabolic adaptations bears great promise for improving treatment efficacy in patients with metastatic diseases (14, 19).
Our laboratory has developed an in vitro model of acquired drug resistance based on chronic exposure of HT29 colorectal cancer cells to incrementally increasing doses of SN38, oxaliplatin, or 5-FU (20, 21). The selected resistant cells maintain a stable chemoresistant phenotype and provide an opportunity to study mechanisms of single-agent resistance. Our unbiased proteomic profiling studies comparing parental cells with resistant cells showed that many metabolic enzymes involved in mitochondrial respiration, glycolysis, and lipogenesis are altered (22). ATP citrate lyase (ACLy), the first-step rate-limiting enzyme for de novo lipogenesis, is one of the proteins that are upregulated in the resistant colorectal cancer cells. Recently, ACLy has been investigated as an anti-cancer therapeutic target (23); however, the contribution of ACLy to drug resistance of cancer cells remains to be elucidated. In this study, we tested the hypothesis that ACLy activation plays a role in the development of drug resistance in colorectal cancer cells. We found that activations of the ACLy and AKT pathways play critical roles in colorectal cancer cell resistance to SN38.
Materials and Methods
Cell lines and in vitro chemoresistance model
The human colorectal cancer cell line HT29 was obtained from the American Type Culture Collection (ATCC). Oxaliplatin- and 5-FU–resistant cell lines were developed in our laboratory as previously described (20, 21). SN38-resistant cell lines were developed by using a similar protocol. Briefly, parental HT29 cells were exposed to an initial SN38 dose of 1 nmol/L and cultured to a confluency of 80% for 3 passages (∼6 weeks). The cells that survived the initial SN38 treatment were then exposed to 5 nmol/L SN38 for 3 passages (∼8 weeks) and to 10 nmol/L for 3 more passages (∼8 weeks). Finally, the SN38 concentration was increased to the clinically relevant plasma drug concentration of 15 nmol/L for 3 passages (∼10 weeks). The surviving resistant cells were named HT29-SNR.
All cells were cultured in minimal essential medium (MEM) containing 5 mmol/L glucose and supplemented with 10% FBS, vitamins, nonessential amino acids, penicillin/streptomycin, sodium pyruvate, and l-glutamine (Life Technologies). HT29-SNR cells were continuously cultured in 15 nmol/L SN38 unless otherwise indicated. Cell viability was measured using a Vi-Cell XR cell viability analyzer (Beckman Coulter). In vitro experiments were carried out at 70% cell confluency at least 3 times. All cell lines were authenticated by short tandem repeat sequencing and matched with 100% accuracy to the ATCC database.
To stably overexpress ACLy in HT29 cells, lentivirus transduction particles were generated using 293 packaging cells transfected by plasmid constructs containing full-length ACLy cDNA. ACLy cDNA was subcloned into the mammalian expression vector pCDNA3.1(−) (Invitrogen) from bacterial cloning vector pOTB6 (MG1813; ATCC) using EcoR1 and HindIII sites and subsequently cloned into a GFP-expressing vector for virus production. Virus particles and polybrene (10 μL/mL) were added to cells at 70% confluency in 6-well plates for 24 hours. After cell expansion for one passage, GFP-positive cells were subjected to fluorescence-activated cell sorting (FACS) to obtain cells stably expressing ACLy.
Flow cytometric analysis for cell death
Apoptotic cells and total dead cells (sub-G1 population) were quantified by Annexin V/propidium iodide (PI) staining and flow cytometry. Apoptosis was analyzed by an Annexin V assay kit (BD Biosciences Pharmingen) according to the manufacturer's instructions. For sub-G1 phase quantification, cells grown in 6-cm-diameter petri dishes were harvested with trypsin and washed in PBS. Cells were then resuspended in 10% ethanol for fixation. Next, 100 μL of a cell suspension (105 cells) was stained with PI at room temperature for at least 15 minutes in the dark. Cells were then analyzed in a FACSCalibur flow cytometer (Becton Dickinson). Data were analyzed by FlowJo software (Tree Star).
RNA interference for ACLy knockdown
SmartPool siRNA oligonucleotides for ACLy and a negative control were purchased from Dharmacon. Four independently validated siRNAs were transfected into HT29-SNR cells together at a final concentration of 40 nmol/L by Lipofectamine (Invitrogen) according to the manufacturer's protocol. First, 120 pmol of siRNA and 16 μL of Lipofectamine were mixed in 500 μL of Opti-MEM medium (10 nmol/L final concentration for each individual siRNA in the SmartPool). After 20 minutes of incubation, the mixture was added to cells at 70% confluency plated in a 6-well plate (2 mL final volume). Fresh medium was added to cells after 6 hours. Twenty-four hours after transfection, cells were either collected for protein harvest or continued for SN38 treatment for additional 48 hours before analysis for apoptosis markers by Western blotting.
Spectrophotometric assay for ACLy activity
ACLy activity was determined as described previously (24) in an assay mixture containing 100 mmol/L Tris-HCl (pH 7.4), 10 mmol/L MgCl2, 20 mmol/L potassium citrate, 1 mmol/L dithiothreitol, 10 U of malic dehydrogenase, 300 μmol/L coenzyme A (CoA), 280 μmol/L NADH, and various amounts of purified ACLy (BPS Bioscience), and optimal conditions for measuring enzyme activities were identified. The reaction was initiated by adding 10 mmol/L ATP at 37°C, and NADH oxidation, as evidenced by a decrease in absorbance at 340 nm, was monitored continuously for 10 minutes using a microplate reader. To measure the activity of GSK165, a small-molecule inhibitor of ACLy (provided by GlaxoSmithKline under a material transfer agreement; 25), 400 μg of purified ACLy was used in a final volume of 100 μl. Controls not containing ACLy were used to account for nonspecific NADH oxidation.
Cell growth inhibition was quantified by MTT assay. First, HT29 cells were seeded in 96-well plates at 3,000 cells in 100 μL per well and incubated overnight. The next day, a 100 μL working stock solution containing GSK165 at 2× final concentration was added to the cell suspension. After 72-hour incubation, 40 μL of the MTT reagent (3 mg/mL) was added to each well, and the mixtures were incubated for 2 to 4 hours. After the supernatant was removed, the formazan precipitates in the cells were dissolved in 100 μL of dimethyl sulfoxide (DMSO). Absorbance at 570 nm was determined using a MultiSkan plate reader (LabSystems). Fractional survival was plotted against the logarithm of the drug dose, and 50% inhibitory concentrations (IC50) were calculated by Prism software (GraphPad Software).
Antibodies against ACLy, p-ACLy (ACLy phosphorylated at the S454 site), AKT, p-AKT S473, cleaved PARP, and cleaved caspase-3 were purchased from Cell Signaling. β-Actin antibody was purchased from Santa Cruz Biotechnology. Whole-cell lysate was collected from cells cultured at 70% confluency before standard Western blot analysis was conducted.
Drugs and other reagents
Oxaliplatin, 5-FU, and SN38 were obtained from The University of Texas MD Anderson Cancer Center pharmacy. Stock drugs of GSK165, phosphoinositide 3-kinase (PI3K) inhibitor wortmannin (Sigma), AKT-specific inhibitor MK2206 (Selleckbio), and fatty acid synthase inhibitor Cerulenin (Sigma) were reconstituted in DMSO at 10 mmol/L and stored in aliquots at −20°C.
For all in vitro experiments, statistical analyses were conducted using the Student t test (Prism). All statistical tests were 2-sided, and P ≤ 0.05 was considered to be significant.
ACLy expression in human colorectal cancer
We searched the Oncomine database using criteria “ACLy, colorectal cancer, normal vs cancer” and found that a total of 18 studies deposited gene expression data on ACLy expression to the database. As summarized in Fig. 1A, the ACLy gene is among the top 1% to 10% overexpressed genes in colon adenocarcinoma compared with normal colon mucosa. The ACLy mRNA level is increased by 2-fold in malignant colon carcinoma tissues according to Oncomine data from 18 independent studies. In addition, our unbiased proteomic profiling suggests that drug-resistant colorectal cancer cells have elevated ACLy expression (22).
Overexpression of ACLy induces chemoresistance in chemo-naïve HT29 cells
To determine the role of ACLy in drug resistance, we stably overexpressed ACLy protein by lentivirus transduction in HT29 cells (Fig. 1B), which have relatively low levels of basal ACLy and p-ACLy expression compared with other colorectal cancer cell lines (Supplementary Fig. S2). Overexpression of ACLy decreased the sensitivity of HT29 cells to SN38, as evidenced by decreased levels of cleaved PARP, an apoptosis marker, on Western blotting (Fig. 1C). Furthermore, PI staining and flow cytometry showed that ACLy overexpression inhibited cell killing by SN38. Figure 1D shows that the sub-G1 (apoptotic and necrotic) cell population decreased from 46.2% ± 2.5% (SEM throughout) to 5.9% ± 0.5% (P < 0.05) after ACLy overexpression following SN38 treatment but to a less extent from 7.5% ± 0.5% to 2.8% ± 0.5% (P < 0.05) following oxaliplatin treatment. We did not observe any protective effect of ACLy overexpression against 5-FU treatment (data not shown). Because the most striking effect was observed with SN38, we conducted further experiments with SN38-treated cells.
Inhibition of ACLy sensitizes chemo-naïve HT29 cells to SN38 treatment
We next explored the relationship between SN38 and ACLy activity, and the effect of targeting ACLy in combination with SN38 treatment on chemo-naïve HT29 cells. Surprisingly, SN38 activated both AKT and ACLy (Fig. 2A). A transient transfection of ACLy siRNA efficiently knocked down ACLy protein for 24 to 72 hours. After a 24-hour ACLy knockdown, we exposed the cells to 20 nmol/L SN38 for an additional 48 hours. Knockdown of ACLy sensitized HT29 cells to SN38 without affecting AKT activity, as evidenced by increased PARP and caspase-3 cleavage (Fig. 2A) and enhanced cell growth-inhibitory effect of SN38 by MTT assay (Fig. 2B).
We next investigated the effect of GSK165, which is a novel ACLy inhibitor modified from 2-hydroxy-N-phenylbenzenesulfonamide pharmacophore, structurally different from the known ACLy inhibitor SB-20499 and its prodrug SB-201076 (24, 25). As shown in Fig. 2C and D, GSK165 inhibited ACLy activity in a concentration-dependent manner and had a growth-inhibitory effect on cancer cells as a single agent with an IC50 of about 30 μmol/L. When 40 μmol/L GSK165 was combined with SN38 treatment, GSK165 sensitized the cells to SN38, as shown by Western blot analysis of PARP and caspase-3 cleavage (Fig. 2E). The chemosensitization effect of GSK165 could be inhibited by overexpression of fatty acid synthase (FASN; Supplementary Fig. S3), suggesting that ACLy-dependent de novo lipogenesis is involved in ACLy-mediated chemoresistance of HT29 cells to SN38.
ACLy activation is associated with acquired resistance to SN38
To investigate whether ACLy is also involved in the development of acquired resistance to SN38, we determined the levels of total ACLy and p-ACLy in chemo-naïve and SN38-resistant HT29 cells (HT29-SNR). We found that the level of p-ACLy, but not total ACLy, was significantly higher in HT29-SNR cells than in parental cells (Fig. 3A). Increased p-ACLy activation may lead to increased de novo lipogenesis. We observed that the numbers of cytoplasmic lipid droplets by electron microscope imaging also increased significantly in drug-resistant cells (Supplementary Materials and Fig. S4). To further confirm the increase of lipid content in HT29-SNR cells is due to de novo lipogenesis, we examined the FASN level and discovered that the HT29-SNR cells significantly upregulated FASN expression compared with the parental HT29 cells (Supplementary Fig. S5A). Treatment with SN38 acutely increased intracellular lipid content in the parental HT29 cells as measured by AdipoRed assay, and this increase can be blocked by a specific FASN inhibitor, cerulenin (Supplementary Materials and Fig. S5B and S5C). Similarly, cerulenin also significantly reduced the lipid content of HT29SNR cells (Supplementary Fig. S5D). These data indicate that SN38-induced intracellular lipid accumulation is due to enhanced de novo lipogenesis.
These findings prompted us to test the possibility that targeting ACLy may resensitize HT29-SNR cells to SN38. However, treatment of cells with GSK165 or with ACLy siRNA failed to sensitize HT29-SNR cells to SN38, as shown by analysis for apoptotic cells by Annexin V–coupled flow cytometry (Fig. 3B) and Western blot analysis for cleaved PARP (Fig. 3C). These data suggest that, in addition to the activation of ACLy, SN38 resistant cells must have acquired other survival mechanisms.
Dual inhibition of ACLy and AKT pathways resensitized HT29-SNR cells to SN38
To explore the mechanisms behind the failure of resensitizing HT29-SNR cells to SN38 by ACLy inhibition, we measured the activity of AKT in response to ACLy knockdown. Knockdown of ACLy activated AKT in HT29-SNR cells regardless of SN38 (Fig. 4A). These data suggest that the activated AKT pathway likely contributes to the failure of resensitizing HT29-SNR cells to SN38 by ACLy inhibition.
Speculating that a dual targeting of the ACLy and AKT pathways may resensitize HT29-SNR cells to SN38, we next tested the effect of a triple combination composed of the PI3K inhibitor wortmannin, GSK165, and SN38 on HT29-SNR cells. The triple combination suppressed activation of both the ACLy and the AKT pathways and successfully resensitized HT29-SNR cells to SN38-induced apoptosis, as shown by increased levels of cleaved PARP and cleaved caspase-3 (Fig. 4B). We next confirmed the role of AKT in chemoresistance of HT29-SNR cells using a more potent AKT inhibitor, MK2206 (26). MK2206 significantly resensitized HT29-SNR cells to SN38 treatment under the condition of ACLy knockdown by siRNA treatment (Fig. 4C). We next treated HT29-SNR cells with a triple combination of MK2206 (2.5 μmol/L, sufficient to suppress AKT activation and ACLy (Supplementary Fig. S6), SN38 (20 nmol/L), and GSK165 (5 μmol/L, concentration reduced to show a better combination effect). Synergistic effects of growth inhibition and resensitization of HT29-SNR cells to SN38-induced apoptosis were observed using the triple combination (Figs. 4D and Supplementary Fig. S6). Figure 5 summarizes the proposed negative feedback regulation of ACLy-AKT and activation of AKT is a compensatory survival pathway under ACLy inhibition in SN38-resistant colorectal cancer cells.
Approximately 50,000 patients with metastatic colorectal cancer die each year in the United States due to resistance to chemotherapy. An understanding of the mechanisms of resistance to individual drugs is critical to develop more effective treatment strategies. Irinotecan is commonly used as part of the first-line chemotherapy backbone for patients with metastatic colorectal cancer. Chemoresistance in cancer cells is known to be mediated by one or more of the following mechanisms: increased drug efflux, enhanced drug inactivation, enhanced DNA damage repair, mutated survival-related genes, deregulated growth factor signaling pathways, increased expression of anti-apoptotic genes, and/or activated intracellular survival signaling following chemotherapeutic stress (9, 10, 12, 27, 28). However, little is known about the role of metabolic changes in drug-resistant cancer cells. Our previous findings indicated that metabolic changes are critically involved in the development of cross-chemoresistance in colorectal cancer cells (29). This study revealed a novel mechanism specific to SN38 resistance that involves activation of ACLy and a potential negative feedback loop in which suppression of ACLy activates AKT signaling.
ACLy is the major enzyme for producing nonmitochondrial acetyl-CoA that is needed for de novo lipogenesis and substrate acetylation (24, 30). Upregulation of ACLy has been found in several types of cancer (31–36). The mechanism by which ACLy is involved in the survival machinery of cancer cells might be complex. Together with the other 2 critical lipogenic enzymes, acetyl-CoA carboxylase and fatty acid synthase, ACLy can promote cell growth and survival by increasing de novo synthesized fatty acids on which cancer cells depend (37). ACLy can participate in epigenetic events of cancer cells by regulating the levels of the key substrate of histone acetyltransferase, acetyl-CoA (30). ACLy can regulate cell survival through coordination with cellular prosurvival metabolic network by controlling cytosolic levels of citrate and oxaloacetate, that is, decrease of cytosolic citrate levels can promote glycolsis by activating phosphofructose kinase (31, 38) and increase of oxaloacetate levels can facilitate gluconeogenesis and glycolysis.
Enhanced de novo lipogenesis is one of the major metabolic alterations used by cancer cells to sustain survival and growth (37). Almost all kinds of carbohydrate can be sources of lipid synthesis, directly or indirectly. As one of the direct substrates (not carbon source) for cytoplasmic fatty acid synthase, acetyl-CoA is mainly derived from the citrate effluxes from the mitochondria, however studies have shown that colonic epithelial cells may have alternative pathways of synthesizing fatty acids independent of ACLy (39), which is likely to be catalyzed by cytosolic acetyl-CoA synthase using acetate as its substrate (40). Our data suggest that ACLy-dependent de novo lipogenesis plays a critical role in SN38 resistance. Several protein kinases have been found to activate ACLy by phosphorylation, which include the nucleotide diphosphate kinase (41), the cAMP-dependent protein kinase (38), and the AKT (42). The concurrent activations of AKT and ACLy by SN38 in the chemo-naïve cells (Fig. 2A) suggest that AKT plays a major role in SN38-induced ACLy activation, which is supported by the data that inhibition of AKT almost completely abolished the phosphorylation of ACLy at its S454 (Supplementary Fig. S6).
Cells commonly respond to growth factor signaling (to regulate cellular metabolism) through activation of AKT, which subsequently activates ACLy (42). The reported indispensability of ACLy in AKT-induced cell transformation of several cancer types, including colorectal cancer, suggests that ACLy is a critical component of the AKT pathway (43). This can be partially explained by the fact that ACLy activity connects 2 major cellular metabolic pathways, namely the glycolysis and the fatty acid synthesis, both of which are regulated by AKT-dependent mechanisms (31, 44). The relationship between the ACLy and AKT pathways seems to be a critical axis on which colorectal cancer cells rely to develop resistance to SN38. However, after achieving the irreversible resistance to SN38, a new compensatory survival relationship between ACLy and AKT is established, that is, suppression of ACLy activity activates AKT (Fig. 5). According to this perspective, inhibition of ACLy releases the “brake” and results in compensatory AKT overactivation, which provides a survival advantage to cells to resist SN38. It has previously been shown that ACLy inhibition leads to a decrease in AKT activity in a chemo-naïve lung cancer cell line (45). The activation of AKT by suppression of ACLy in the SN38-resistant cells is intriguing. The absence of this pathway in the chemo-naïve cells (Fig. 2A) indicates that this feedback loop is unlikely mediated by possible preexisting mechanisms that can link ACLy functions with the AKT pathway in chemo-naïve cells. Considering that SN38 causes genotoxic stress by inducing single-strand and double-strand DNA breakage (8), ACLy can participate in signaling transduction and epigenetic alterations through protein acetylation (30, 43), and the irreversible drug-resistant phenotype of HT29SNR cells, we speculate that new factors emerged from irreversible epigenetic alterations under long-term SN38 exposure mediate the negative feedback loop from ACLy to AKT.
While the underlining mechanism by which suppression of ACLy activates the AKT pathway requires further investigations, the compensatory role of AKT in sustaining cell survival of SN38-resistant cells under conditions of ACLy inhibition can be explained from a metabolic point of view. The effectiveness of co-targeting ACLy and AKT in resensitizing SN38-resistant cells suggests that shared targets by ACLy and AKT play important roles in the development of SN38 resistance of colorectal cancer cells. Previously, we have reported that enhanced aerobic glycolysis is a pivotal factor in the development of cross-drug resistance of colorectal cancer cells (29). Both ACLy and AKT regulate glycolysis. ACLy can enhance glycolysis by activating phosphofructokinase via reducing the cytosolic citrate levels (31). AKT activates glycolysis through multiple molecular mechanisms (46).
Metabolic reprogram in adaptation to chemotherapeutic stresses may be a fundamental mechanism of drug resistance, although cancer cells may achieve some common metabolic alterations, such as elevations of glycolysis and lipogenesis, via different mechanisms in response to different chemotherapeutic reagents. A better understanding of the mechanisms by which cancer cells gain resistance to commonly used chemotherapeutic reagents through metabolic adaptation bears a great potential of identifying novel targets for preventing and overcoming chemoresistance.
Disclosure of Potential Conflicts of Interest
No potential conflicts of interest were disclosed.
Conception and design: Y. Zhou, L.M. Ellis, Z. Weihua
Development of methodology: Y. Zhou, L.R. Bollu, F. Tozzi, X. Ye, R. Bhattacharya, G. Gao, L. Xia, F. Fan
Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): Y. Zhou, L.R. Bollu, F. Tozzi, G. Gao, E. Dupre, J. Lu, S. Bellister
Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): Y. Zhou, F. Tozzi, G. Gao, S. Bellister, Z. Weihua
Writing, review, and/or revision of the manuscript: Y. Zhou, S. Bellister, L.M. Ellis, Z. Weihua
Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): Y. Zhou, G. Gao, L. Xia, J. Lu, S. Bellister
Study supervision: Y. Zhou, L.M. Ellis, Z. Weihua
The authors thank Kenneth Dunner Jr and Robert Langley, PhD, of the Electron Microscope Core Facility of MD Anderson Cancer Center for their assistance in lipid droplet morphology analysis. They also thank Li Huang of the Subcloning Core of the Department of Cancer Biology for generation of lentivirus particles containing the ACLy gene, the Flow Cytometry Core Facility for the Annexin V/PI assay analysis of apoptosis, and Rita Hernandez of the Departments of Surgical Oncology for editorial assistance.
This research was supported, in part, by the NIH (Cancer Center Support Grant CA016672) to L.M. Ellis. F. Tozzi was supported by NIH grant T32 CA009599. Z. Weihua was supported by grants from the American Cancer Society and Congressional Directed Medical Research Programs of the Department of Defense. L. M. Ellis was also supported by the William C. Liedtke Jr, Chair in Cancer Research.
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