A Selective Novel Peroxisome Proliferator-Activated Receptor (PPAR)-α Antagonist Induces Apoptosis and Inhibits Proliferation of CLL Cells In Vitro and In Vivo
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Tumor-specific metabolic changes can reveal new therapeutic targets. Our findings implicate a supporting role for fatty acid metabolism in chronic lymphocytic leukemia (CLL) cell survival. Peroxisome proliferator-activated receptor (PPAR)-α, a major transcriptional regulator of fatty acid oxidation, was recently shown to be upregulated in CLL. To evaluate PPARα as a potential therapeutic target, we developed a highly selective, potent small molecule antagonist of PPARα, NXT629. NXT629 inhibited agonist-induced transcription of PPARα-regulated genes, demonstrating target engagement in CLL cells. Furthermore, NXT629 induced apoptosis of CLL cells even in the presence of a protective microenvironment. To mimic the proliferative lymphoid compartment of CLL, we examined the activity of NXT629 on CLL cells that were stimulated to proliferate in vitro. NXT629 reduced the number of leukemia cells undergoing cell division. In addition, in two xenograft mouse models of CLL (one a model for nondividing and one for dividing CLL), NXT629 reduced the number of viable CLL cells in vivo. Overall, these results suggest that fatty acid metabolism promotes survival and proliferation of primary CLL cells and that inhibiting PPARα gene regulation could be a new therapeutic approach to treating CLL.
Chronic lymphocytic leukemia (CLL) is the most common adult leukemia in the Western world, leading to ∼5,000 deaths annually (1). CLL is characterized by an accumulation of monoclonal mature B cells in blood, secondary lymphoid tissues and bone marrow. Despite major advances in the field, there is no curative therapy for CLL to date (2). Current treatment approaches aim at achieving minimal residual disease, which is associated with superior long-term outcome (3). Treatment avenues, such as those targeting pathways downstream of the B cell receptor, such as Syk, Btk and PI3Kδ (4, 5, 6, 7, 8, 9), that are currently being evaluated in clinical trials or have recently been approved by the U.S. Food and Drug Administration (such as ibrutinib) are focused on kinases. Surprisingly, little attention has been given to targeting metabolic enzymes in CLL. While the traditional view has been that cancer cells are fueled by glucose, named the “Warburg effect” after Otto Warburg (10,11), recent studies demonstrate the involvement of fatty acid oxidation (FAO) in cancer cell viability. Some solid tumors including prostate, ovarian and renal cell carcinoma rely on fatty acids to satisfy their metabolic needs (12, 13, 14, 15, 16). Solid tumors that are initially dependent on glucose can undergo a metabolic switch upon detachment from the extracellular matrix and start depending on FAO for survival (12). In addition, hypoxia and oncogenic RAS increase fatty acid uptake by tumor cells (17). Inhibition of FAO with Etomoxir, an irreversible small molecule inhibitor of CPT1A, the rate-limiting enzyme for fatty acid import into mitochondria (18), sensitized AML cells to apoptosis induction by ABT-737, an inhibitor of Bcl-2 and Bcl-xL (19). Furthermore, some evidence points toward a critical role for FAO in the viability of leukemia-initiating cells. For instance, the CPT1A inhibitor Etomoxir decreases the number of leukemia-initiating cells in primary human AML samples (19), and FAO signaling downstream of PML was found to be critical for maintenance and function of hematopoietic stem cells and possibly leukemia-initiating cells (20).
Peroxisome proliferator-activated receptors (PPARs) are a family of ligand activated nuclear hormone receptors comprised of three isoforms: PPARα, PPARδ and PPARγ. PPARα, also known as NR1C1 (nuclear receptor subfamilyl, group C, member 1) is a major transcriptional regulator of lipid metabolism. Endogenous PPARα ligands include free fatty acids and eicosanoids as well as oleoylethanolamide (OEA), a naturally occurring lipid (21). Upon ligand binding, PPARα induces transcription of a number of genes, resulting in a shift toward β-oxidation (for example, CPT1A) and away from glucose oxidation (for example, PDK4) (22).
It was recently shown that compared with normal B cells, CLL cells overexpress PPARα, rendering them dependent on β-oxidation for energy (23). This metabolic strategy helps account for some of the pathogenic characteristics of CLL, including immunosuppression and drug resistance. These reports establish PPARα as a promising molecular target for the treatment of this cancer. However, to date, no clinically relevant, selective PPARα antagonists have been available to address this idea in human trials. We have recently described the discovery and synthesis of a family of novel, selective and bioavailable PPARα antagonists that can bind reversibly to the ligand binding site and effectively compete with both synthetic and endogenous PPARα agonists (24). Herein, we evaluated the activity of the PPARα antagonist NXT629 on CLL viability and proliferation in vitro as well as tumor burden in vivo.
Materials and Methods
Patient Samples, Isolation of CLL B cells, Cell Culture and Reagents
Blood samples were collected from patients at The Feinstein Institute for Medical Research, North Shore-LIJ Health System (Manhasset, NY, USA), and the Sunnybrook Odette Cancer Center (Toronto, Canada) who satisfied diagnostic and immunophenotypic criteria for common B-cell CLL after providing written informed consent in compliance with the Declaration of Helsinki (25) and the Institutional Review Board of the North Shore-LIJ Health System and the Sunnybrook Health Sciences Center. Blood was collected from patients for whom clinical information and laboratory data were available. For a set of patients (from N Chiorazzi), immunoglobulin heavy-chain variable region gene (IGHV) and immunoglobulin light-chain variable region gene (IGLV) DNA sequences were also available. For some experiments, the CLL cells were purified from frozen peripheral blood mononuclear cells (PBMCs) via negative selection by using anti-CD2 and anti-CD14 magnetic beads (Miltenyi Biotech).
Isolation of B Cells from Healthy Volunteers
Peripheral blood mononuclear cells were isolated from the blood of normal volunteers over a Ficoll-Hypaque density gradient. Anonymous blood samples were purchased from the San Diego Blood Bank; therefore, no Institutional Review Board approvals were necessary. B cells were isolated by positive selection using CD19+ beads (Milenyi Biotech) per the manufacturer’s instructions. Cells were cultured and viability was monitored as described below.
Measurement of Cell Viability
CLL cells were cultured at 2 × 105 cells/mL in 100 µL media in 96-well plates (Costar, Corning Inc.) in RPMI 1640/10% fetal calf serum (FCS). All inhibitors were prepared in dimethyl sulfoxide (DMSO) (0.1% final concentration), which was used as vehicle control in all experiments. Determination of CLL cell viability was based on the analysis of mitochondrial transmembrane potential (Δψ)m) using 3,3′-dihexyloxacarbocyanine iodide (DiOC6) (Invitrogen), and cell membrane permeability to propidium iodide (PI) (Sigma-Aldrich). For viability assays, 100 µL of the cell culture was collected at the indicated days and mixed with media containing 100 µL of 40 µmol/L DiOC6 and 10 µg/mL PI. The cells were then incubated at 37°C for 15 min and analyzed within 30 min by flow cytometry by using the flow cytometer Accuri C6 (Accuri). Data were analyzed by using the CFlowPlus software (Accuri). The percentage of viable cells was determined by gating on PI negative and DiOC6 bright cells.
Macrophages (J774A.1) were obtained from ATCC (ATCC® TIB67™ and CRL-11882™) and plated in 96-well plates at a density of 50,000 cells per well. OP9 cells were also obtained from ATCC (ATCC® CRL-2749™) and plated at 10,000 cells/well. Macrophages and OP9 cells (26) were exposed to 10 µg mitomycin C (Sigma-Aldrich M4287) for 3 h to prevent proliferation. The cells were then washed three times with media to remove the mitomycin C. Cells were plated in RPMI/10% FCS, and CLL cells were subsequently plated over the layer of mitomycin C-treated macrophages or OP9 cells. NXT629 or vehicle control was added to the cells at the beginning of the coculture and incubated in a 37°C/5% CO2 incubator for 7 d. After 7 d, the CLL cells were collected and stained with DiOC6/PI and analyzed for viability as described above.
CLL Proliferation Assay
PBMCs were isolated from the blood of normal volunteers (purchased anonymously from the San Diego Blood Bank) over a Ficoll-Paque PLUS density gradient (GE Healthcare). CD2+ T cells were isolated from PBMCs by positive selection using anti-CD2 beads (Miltenyi Biotech) following the manufacturer’s instructions. To activate T cells, they were cultured on a 24-well plate (Costar/Corning Inc.) at 1 × 106 cells/mL in 1 mL of culture media (RPMI 1640 supplemented with 10 mmol/L HEPES [GIBCO-BRL], penicillin [100 U/mL]-streptomycin [100 µg/mL] (Gibco/Invitrogen) and 10% FCS [ATCC]) in the presence of 20 µL CD3/CD38 Dynabeads (Invitrogen) per 106 T cells for 3 d. T cells were collected, resuspended at 2 × 107 cells/mL in fresh medium containing 12 µg/mL mitomycin C and incubated for 3 h at 37°C. At this point, the T cells were washed four times with fresh media and frozen by using Recovery Cell Culture freezing media (Gibco/Life Technologies) in liquid N2 until use. CLL proliferation induced by frozen versus fresh T cells was comparable (data not shown). Thus, frozen T cells were used for all experiments.
To set up the CLL proliferation assay, CLL PBMCs were thawed from frozen vials and cultured at 1.4 × 105 CLL cells/well in 100 µL of the above-listed culture media for 2 h in the presence of PPARα antagonist or vehicle control. Mitomycin C-treated, activated T cells were added at 6 × 104 T cells/well in 100 µL of the above-listed culture media supplemented with hrIL-4 (5 ng/mL, R&D Systems) and hrIL-10 (15 ng/mL, R&D Systems). Cells were cultured for 5–8 d. In some experiments, CLL cells were prelabeled with carboxyfluoresceinsuccinimidyl ester (CFSE) (Invitrogen) after the manufacturer’s instructions before coculture. Five days after coculture, CLL cells were stained with CD19-APC (Becton Dickinson), and proliferation was assessed by flow cytometry (Accuri cytometer) gating on CD19+ CFSE+ cells. In the majority of the experiments, if not otherwise indicated, the number of viable CLL cells was analyzed by staining with DiOC6/PI as described above and collecting each sample for 30 s via flow cytometry. The number of viable cells was determined by gating on DiOC6 bright and PI negative cells.
Cell Cycle Analysis
CLL cells were cultures as above to trigger proliferation. At d 8, cells were collected and analyzed for cell cycle progression by using the FxCycle PI/RNase staining solution (Molecular Probes, Life Technologies) according to the manufacturer’s instructions.
Isolation of RNA and cDNA Synthesis
Purified CLL cells were plated at 6.8 × 106 cells/mL in 1 mL media in a 12-well plate. Cells were exposed to antagonist NXT629 for 2 h, followed by an agonist for 48 h, as described in the respective figure legends. RNA was isolated from CLL cells by using the RNeasy kit (Qiagen). A total of 100 ng RNA was used in each cDNA reaction by using the IScript reaction mix (Bio-Rad) following the manufacturer’s instructions.
Real-Time Reverse Transcriptase Polymerase Chain Reaction
Reverse transcription reactions were performed by using the iTaqUniversal SYBR® Green supermix (Bio-Rad) following the manufacturer’s instructions. Each reaction contained 2.5 ng reverse-transcribed RNA (based on the initial RNA concentration) in 20 µL final reaction volume. The reaction conditions were as follows: 95°C for 30 s, 45 cycles of 95°C for 4 min followed by 59°C for 5 min, 65°C for 5 min and finally 95°C for 5 min. Primer sequences used were as follows: β-actin-F: GCT GTG CTA CGT CGC CCT G, β-actin-R: GGA GGA GCT GGA AGC AGC C, PDK4-F: GGAGC ATTTCTCGCGCTACA, and PDK4-R: ACAGGCAATTCTTGTCGCAAA. Primers were synthesized by Integrated DNA Technologies. The polymerase chain reaction (PCR) was carried out by using the C1000Touch Thermal Cycler (Bio-Rad).
CLL Mouse Model
Two different CLL mouse models were evaluated. NOD/Shi-scid,ycnull (NSG) mice, a NOD/SCID-derived strain, that lacks the IL-2 receptor family common cytokine receptor γ chain gene (μc), rendering animals completely deficient in lymphocytes, including natural killer cells, were used for this study. Female NSG mice (12–14 wks at study initiation) were purchased from The Jackson Laboratory. Animals were given food and water ad libitum and allowed to acclimate for at least 1 wk before initiation of experiments. All protocols were approved by the Inception Sciences Institutional Animal Care and Use Committee.
Model for resting CLL. CLL PBMCs from two patients were pooled, and CFSE was labeled and randomized among the groups. The 108 CFSE-labeled cells were delivered by an intravenous bolus injection (50 µL) into the tail vein of NSG mice. Immediately after injection of CLL cells, groups of five mice received daily dosing of vehicle control (saline, 10 mL/kg, intraperitoneal), NXT629 at 30 mg/kg or fludarabine at 50 mg/kg. Mice were sacrificed 4 wks after engraftment, and the splenocytes were stained with hCD19 and hCD5 and analyzed by flow cytometry.
Model for proliferative CLL. The Institutional Review Board and the Institutional Animal Care and Utilization Committee of the North Shore-LIJ Health System sanctioned these studies. T cells were purified from CLL PBMCs using Milteny anti-CD3 beads, resuspended in 1 × 106 cells/mL and stimulated with anti-CD3/CD28 Dynabeads (30 µL/mL) in the presence of IL-2 (36 U/mL) in RPMI 1640/10% FCS for 3 d. Next, beads were removed from the cultures, and the cells were cultured in media supplemented with IL-2 for an additional 4 d. Preactivated human T cells (5 × 105) were administered in 4- to 8-wk-old NSG mice (The Jackson Laboratory) by injection into the retro-orbital plexus (50 µL). After confirming the presence of human T cells in the blood of recipient mice (10 d after injection), CLL PBMCs from the same patient (2 × 107) were delivered by an intravenous (50 µL) injection into the retroorbital plexus. At the time of CLL cell injection, mice received vehicle control or NXT629, 30 mg/kg of mouse weight, which was given by intraperitoneal injections daily for 2 wks. All mice were killed at the end of experiment, and the spleen and bone marrow (BM) were collected for flow cytometric analyses. Spleen and BM cells were stained by using anti-mCD45, anti-hCD45, anti-hCD5, anti-hCD19, anti-hCD4 and anti-hCD8 antibodies.
Statistical significance was determined by using the Student t test. The p values <0.05 were considered significant. Median inhibitory concentration (IC50) values were determined using nonlinear regression (curve fit) analysis with Prism software (GraphPad Software).
All supplementary materials are available online at https://doi.org/www.molmed.org .
NXT629 Inhibits Transcription of PPARα Target Genes
IC50 values (inhibition in antagonist mode) for human PPARα, PPARδ and PPARγ.
IC50 values µmol/L using NXT629 (inhibition in antagonist mode) for human nuclear hormone receptors?
0.077 ± 35
6.0 ± 3.2
15 ± 19
12 ± 8
26 ± 21
57 ± 27
PPARα Antagonist Is Cytotoxic to CLL Cells Even in the Presence of a Protective Microenvironment
CLL cells in lymphoid organs are in contact with their microenvironment, and cells of the microenvironment can protect CLL cells from spontaneous and drug-induced apoptosis (31). It is therefore critical to evaluate new compounds in the context of the cellular microenvironment. Thus, we examined whether NXT629 could kill CLL cells when cocultured with macrophages, which can protect CLL cells from spontaneous apoptosis in vitro (32). As expected, macrophages protected CLL cells from spontaneous apoptosis in vitro (Figure 2B), and addition of NXT629 to these cocultures caused a significant reduction in CLL cell viability. After 6 d, the majority of CLL cells underwent apoptosis in the presence of 10 µmol/L NXT629 (Figure 2B). Because a frontline therapy for CLL is fludarabine (2), we compared NXT629 and fludarabine for their potential to induce apoptosis of CLL cells in the presence of accessory cells. Under the same coculture conditions, CLL cells were completely protected from fludarabine-induced apoptosis (Figure 2C), at doses that were cytotoxic to CLL cells cultured in the absence of macrophages (Figure 2D), whereas NXT629 induced apoptosis of CLL cells under both conditions (Figures 2A, B). Preadipocytes (OP9 cells ) were evaluated for their ability to protect CLL cells from spontaneous apoptosis, since they play a supportive role in other cancers and could be a potential source of lipids in vivo (33,34). Preadipocytes (OP9 cells) also increased viability of CLL cells, and this effect was completely blocked by using 10 µmol/L NXT629 (Figure 2E).
PPARα Antagonist Inhibits CLL Proliferation
NXT629 Reduces CLL Tumor Burden in Two Adoptive Transfer Mouse Models
Because a fraction of CLL cells proliferate in vivo (35), it is critical to evaluate new treatment modalities on the proliferative compartment. It has been shown that autologous T cells promote CLL cell survival and proliferation in NSG mice (38). To model the proliferative CLL compartment in vivo, a modified protocol of the xenograft model described by Bagnara et al. (38) was applied. In brief, first, in vitro activated CLL T cells were adoptively transferred into NSG mice. Subsequently, autologous CLL B cells were given to those mice in which T-cell engraftment was documented by the presence of CD3+ cells in the blood. This approach is a model of CLL B-cell growth in vivo, since the transferred leukemic cells proliferate extensively, often exceeding six to seven divisions on the basis of CFSE dilution analyses during the course of the study. However, the approach is not a model of CLL disease, since the transferred clone survives for 4–12 wks; this time interval varies on the basis of the CLL sample analyzed. Nevertheless, during this window, the effects of various therapies on CLL growth can be effectively studied. Proliferating CLL B cells are found most often in the spleen and to a lesser extent in the bone marrow. Lymph node infiltration is rarely seen during the time frame of these studies.
Mice treated in this manner were dosed daily with 30 mg/kg NXT629 IP, and 2 wks after B-cell administration, recipients were killed and splenocytes were stained with mCD45/hCD45/hCD19/hCD5. As seen in the other CLL model, a marked reduction in both the percentage and the absolute number of hCD19+/hCD5+ CLL cells was observed in NXT629-treated animals (Figure 4B). These results suggest that NXT629 delays disease progression of CLL in vivo.
Despite major advances in the field, there is no curative therapy for CLL to date (2). All patients inevitably relapse and retreatment is often limited by resistance to chemotherapy. Thus, new therapies are needed. Current treatment approaches aim at achieving minimal residual disease, which is associated with superior long-term outcome (3). A major focus in the field has been on inhibition of kinases, and little attention has been given to metabolic pathways. Recent reports have highlighted the role of PPARα and FAO in cancer (14,19,39, 40, 41). PPARα KO mice completely suppress metastasis and growth of primary tumor in Lewis lung carcinoma and melanoma models >100 d after tumor implantation (42). PPARα is overexpressed in CLL cells and helps to protect them from harsh microenvironmental conditions. MK886, a small molecule that is reported to have PPARα antagonist properties, exhibits anti-CLL activity in vitro and in vivo (23,30). Taken together, these observations made us consider that PPARα may be an important therapeutic target for CLL and other cancers that use FAO as a metabolic strategy. We have also performed in-house experiments showing that, unlike NXT629, MK886 is not selective, since it inhibits the following nuclear hormone receptors: ERβ (7.5 µmol/L), TR (11 µmol/L) and GR (10 µmol/L) at IC50 levels that are lower than or similar to PPARδ (17 µmol/L), PPARγ (14 µmol/L) and PPARα (18 µmol/L). We evaluated MK886 in the luciferase reporter assay and found that its IC50 for inhibition of PPARα-driven luciferase expression overlapped with the IC50 for cellular cytotoxicity in the CHO cells. The strong overlap in toxicity prevents a clear interpretation regarding whether this molecule does indeed inhibit PPARα, leading to a decrease in luciferase signal or that the cytotoxic effect leads to a decrease in luciferase signal. NXT629 does not suffer from the same problem, since it is not cytotoxic in CHO cells and is highly selective for PPARα (100-fold or more selective) (Table 1). To date, there are no highly selective PPARα antagonists that are available for clinical studies. Taken together, these results strongly suggest that MK886 is not a good molecule to study PPARα inhibition, and therefore our study is the first one to show the effect of PPARα inhibition on CLL cells.
The small molecule PPARα antagonist NXT629 induced apoptosis in resting CLL cells and inhibited proliferation of CLL cells in vitro (Figures 2, 3), but was less cytotoxic to B cells isolated from healthy volunteers (Supplementary Figure S1). Because accessory cells can rescue CLL cells from spontaneous and drug-induced apoptosis (43, 44, 45), as well as protect CLL cells from fludarabine-induced apoptosis in vitro (31), it is essential to evaluate potential therapeutics in CLL accessory cell cocultures. Importantly, NXT629 induced apoptosis of CLL cells in the presence of the microenvironment, whereas fludarabine was completely inactive (Figure 2C).
The doses needed to see an effect in CLL experiments are significantly higher than the IC50 value in the luciferase reporter assay. We can only speculate as to why this is the case. The luciferase reporter assay also requires less agonist to induce gene expression and generally appears more sensitive to both PPARα induction and inhibition. This method is an artificial system with CHO cells overexpressing a reporter construct. It is not known which and how many molecules of corepressor and coactivator are present in CHO cells in comparison to CLL cells. All of this could play are role. However, the dose that is needed to engage the target in CLL cells in vitro is also the dose that leads to cell death, indicating that the observed effect on CLL cells in on target.
Furthermore, NXT629 was also found to lower CLL tumor burden in two different CLL mouse models in vivo (Figure 4). One CLL mouse model was adopted from Herman et al. (37). In contrasts to Herman et al., who showed proliferation of CLL cells 3–4 wks postxenograft in the spleen, we did not observe proliferation of CLL cells in the spleen. These differences could relate to technical differences or differences in the patient samples used in our studies. Therefore, our in vivo experiment tested the effect of PPARα inhibition on resting CLL cells, and the decrease in tumor burden is most likely due to cytotoxic effects on the tumor, which is in concordance with the in vitro results. The second tumor model (38), also an adoptive transfer model of CLL, was initially described by Bagnara et al. In this model, CLL cells proliferate because of the presence of activated autologous T cells. In concordance with the in vitro proliferating CLL cultures, NXT629 lowered tumor burden of proliferating CLL cells in vivo (Figure 4B). In this mouse model, the majority of CLL cells reside in the spleen; therefore, the splenocytes were analyzed, and both the percentage as well as the absolute number of CLL B cells were reduced in NXT629-treated animals. In addition to the spleen, bone marrow was also analyzed. Although significant engraftment of CLL cells in bone marrow only occurred in two to three mice per group, the same trend was observed as in the spleen. NXT629-treated mice contained a lower number of CD19+/CD5+ human B cells in the bone marrow (data not shown). Interestingly, trough levels as low as 50 nmol/L NXT629 measured in plasma were sufficient to cause a significant reduction in tumor burden, whereas in vitro, micromolar levels are needed. One explanation for the discrepancy between the active dose in vitro and in vivo could be that CLL cells in their in vivo microenvironment are more dependent on β-oxidation compared with in vitro culture conditions and thus are more sensitive to its inhibition. On the other hand, cell culture media contain high levels of glucose, which could provide one possible alternative energy source to fatty acids in vitro and therefore lower the dependency on FAO and reduce the sensitivity to PPARα inhibition. However, removal of glucose in vitro did not increase sensitivity of CLL cells to PPARα inhibition (data not shown), ruling out the possibility of a metabolic switch to glucose in vitro. Another alternative energy source could be glutamine, whose role remains to be investigated. A further consideration was that CLL cells might encounter hypoxic conditions in vivo and thus be more sensitive to inhibition of PPARα. However, culture of CLL cells in vitro at low oxygen failed to increase sensitivity to PPARα inhibition (data not shown).
Alternatively, NXT629 could affect certain cells of the microenvironment in vivo, which after exposure to NXT629 could withdraw support factors. As a consequence of this, CLL cells could undergo apoptosis. Indeed, it is likely a combination of factors where in vivo the antagonist acts on both the CLL cells and the microenvironment. All these questions remain to be investigated.
Overall, our preclinical data demonstrate sensitivity of CLL cells to PPARα inhibition and show PPARα antagonist-mediated cytotoxicity in vitro and in vivo, suggesting that antagonism of PPARα might be a potent and safe new therapeutic target for CLL. Spaner et al. (23) showed that PPARα expression was highly associated with advanced-stage disease. Thus, PPAR expression could be used to select the patient population that would benefit from treatment. Blocking a fundamental fuel source might not allow for escape mutants as, for example, inhibition of kinases does. However, combination therapies will most likely be more efficacious.
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We would like to thank all employees of Inception Sciences for their support and contribution of this program.
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