Anticancer Activities of Pterostilbene-Isothiocyanate Conjugate in Breast Cancer Cells: Involvement of PPARγ

Trans-3,5-dimethoxy-4′-hydroxystilbene (PTER), a natural dimethylated analog of resveratrol, preferentially induces certain cancer cells to undergo apoptosis and could thus have a role in cancer chemoprevention. Peroxisome proliferator-activated receptor γ (PPARγ), a member of the nuclear receptor superfamily, is a ligand-dependent transcription factor whose activation results in growth arrest and/or apoptosis in a variety of cancer cells. Here we investigated the potential of PTER-isothiocyanate (ITC) conjugate, a novel class of hybrid compound (PTER-ITC) synthesized by appending an ITC moiety to the PTER backbone, to induce apoptotic cell death in hormone-dependent (MCF-7) and -independent (MDA-MB-231) breast cancer cell lines and to elucidate PPARγ involvement in PTER-ITC action. Our results showed that when pre-treated with PPARγ antagonists or PPARγ siRNA, both breast cancer cell lines suppressed PTER-ITC-induced apoptosis, as determined by annexin V/propidium iodide staining and cleaved caspase-9 expression. Furthermore, PTER-ITC significantly increased PPARγ mRNA and protein levels in a dose-dependent manner and modulated expression of PPARγ-related genes in both breast cancer cell lines. This increase in PPARγ activity was prevented by a PPARγ-specific inhibitor, in support of our hypothesis that PTER-ITC can act as a PPARγ activator. PTER-ITC-mediated upregulation of PPARγ was counteracted by co-incubation with p38 MAPK or JNK inhibitors, suggesting involvement of these pathways in PTER-ITC action. Molecular docking analysis further suggested that PTER-ITC interacted with 5 polar and 8 non-polar residues within the PPARγ ligand-binding pocket, which are reported to be critical for its activity. Collectively, our observations suggest potential applications for PTER-ITC in breast cancer prevention and treatment through modulation of the PPARγ activation pathway.


Introduction
The incidence of cancer, in particular breast cancer, continues to be the focus of worldwide attention. Breast cancer is the most frequently occurring cancer and the leading cause of cancer deaths among women, with an estimated 1,383,500 new cases and 458,400 deaths annually [1]. Many treatment options, including surgery, radiation therapy, hormone therapy, chemotherapy, and targeted therapy, are associated with serious side effects [2][3][4][5]. Since cancer cells exhibit deregulation of many cell signaling pathways, treatments using agents that target only one specific pathway usually fail in cancer therapy. Several targets can be modulated simultaneously by a combination of drugs with different modes of action, or using a single drug that modulates several targets of this multifactorial disease [6].
Peroxisome proliferator-activated receptors (PPAR) are ligandbinding transcription factors of the nuclear receptor superfamily, which includes receptors for steroids, thyroids and retinoids [7,8]. Three types of PPAR have been identified (a, b, c), each encoded by distinct genes and expressed differently in many parts of the body [8]. They form heterodimers with the retinoid X receptor, and these complexes subsequently bind to a specific DNA sequence, the peroxisome proliferating response element (PPRE) that is located in the promoter region of PPARc target genes and modulates their transcription [9]. PPARc is expressed strongly in adipose tissue and is a master regulator of adipocyte differentiation [10]. In addition to its role in adipogenesis, PPARc is an important transcriptional regulator of glucose and lipid metabolism, and is implicated in the regulation of insulin sensitivity, atherosclerosis, and inflammation [10,11]. PPARc is also expressed in tissues such as breast, colon, lung, ovary, prostate and thyroid, where it regulates cell proliferation, differentiation, and apoptosis [12][13][14].
Although it remains unclear whether PPAR are oncogenes or tumor suppressors, research has focused on this receptor because of its involvement in various metabolic disorders associated with cancer risk [15][16][17]. The anti-proliferative effect of PPARc is reported in various cancer cell lines including breast [18][19][20][21], colon [22], prostate [23] and non-small cell lung cancer [24]. Ligand-induced PPARc activation can induce apoptosis in breast [13,20,25,26], prostate [23] and non-small cell lung cancer [24], and PPARc ligand activation is reported to inhibit breast cancer cell invasion and metastasis [27,28]. Results of many studies and clinical trials have raised questions regarding the role of PPARc in anticancer therapies, since its ligands involve both PPARcdependent and -independent pathways for their action [29].
Previous studies showed that thiazolidinediones can inhibit proliferation and induce differentiation-like changes in breast cancer cell lines both in vitro and in xenografted nude mice [13,30]. Alternately, Abe et al. showed that troglitazone, a PPARc ligand, can inhibit KU812 leukemia cell growth independently of PPARc involvement [31]. In addition to in vitro studies, in vivo administration of PPARc ligands also produced varying results. The use of troglitazone was reported to inhibit MCF-7 tumor growth in triple-negative immunodeficient mice [13] and in DMBA-induced mammary tumorigenesis [32], and administration of a PPARc ligand (GW7845) also inhibited development of carcinogen-induced breast cancer in rats [33]. In contrast, a study by Lefebvre et al. showed that PPARc ligands, including troglitazone and BRL-49653, promoted colon tumor development in C57BL/6JAPCMin/+ mice, raising the possibility that PPARc acts as a collaborative oncogene in certain circumstances [34]. It thus appears that PPARc activation or inhibition can have distinct roles in tumorigenesis, depending on the cancer model examined. Hence determining possible crosstalk between PPARc and its ligand in cancer is critical for the development of more effective therapy.
Trans-3,5-dimethoxy-4-hydroxystilbene (PTER) is an antioxidant found primarily in blueberries. This naturally occurring dimethyl ether analog of resveratrol has higher oral bioavailability and enhanced potency than resveratrol [35]. Based on its antineoplastic properties in several common malignancies, studies suggest that PTER has the hallmark characteristics of an effective anticancer agent [36][37][38][39][40]. Recent research from our laboratory showed that PTER-ITC conjugate (Fig. 1A), a novel class of hybrid compound synthesized by appending an isothiocyanate moiety to the PTER backbone, can induce greater cytotoxicity in tumor cells than PTER alone [41,42]. In human breast and prostate carcinoma cells, PTER-ITC induces strong anticancer activity at a much lower dose than the PTER parent compound [41,42].
Here we analyzed the anti-cancer activity of PTER-ITC in MCF-7 and MDA-MB-231 breast cancer cells. As PPARc mediates anti-tumor activity in a variety of cancer types, we hypothesized that PTER-ITC could modulate the activity of PPARc pathway in breast cancer cells and inhibit tumor cell growth. Our results show that PTER-ITC induced apoptosis in breast cancer cells through caspase activation, which increased the Bax/Bcl-2 ratio and downregulated survivin. Our molecular docking study also demonstrated that PTER-ITC make contact with amino acids within the ligand-binding pocket of PPARc that are crucial for its activation. We found that PPARc activation has an important role in PTER-ITC-induced apoptosis and reduced survivin levels. Our studies thus provide evidence for the usefulness of PTER-ITC in breast cancer therapy involving various pathways, including PPARc.

Cell lines and culture
Three breast cancer cell lines (MCF-7, MDA-MB-231 and T47D) with distinct characteristics were obtained from the National Center for Cell Science (NCCS; Pune, India). MCF-7 and T47D are estrogen receptor (ER)-positive and lack HER-2 expression, while MDA-MB-231 is ER-negative and has low HER-2 expression. MCF-7 cells express wild-type p53, whereas MDA-MB-231 and T47D express mutant p53. All three cell lines express PPARc protein. T47D cells were maintained in RPMI medium supplemented with 2 mM L-glutamine, 4.5 g/L glucose and 0.2 U/ml insulin. MCF-7 and MDA-MB-231cells were maintained in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum (heat-inactivated) (both from Invitrogen, Life Technologies) and 1% antibiotic mix (100 U/ml penicillin, 100 mg/ml streptomycin) at 37uC, 5% CO 2 in a humidified atmosphere.

Cytotoxicity assays
The anti-proliferative effect of PTER and PTER-ITC was determined by the MTT assay as described [41]. Briefly, MCF-7 and MDA-MB-231 cells were seeded at a density of ,56103 cells/well in a 96-well microtiter plate and incubated overnight. Cells were then exposed to increasing PTER and PTER-ITC concentrations (1, 10, 20, 40 and 60 mM) for 24 h. Control cells were treated with 0.1% DMSO (vehicle control). The effect of the inhibitor GW9662 on PTER-ITC-induced cell death was also studied to evaluate involvement of PPARc activation in this process. After 24 h, cultures were assayed by addition of 20 ml MTT (5 mg/ml) and incubation (4 h, 37uC). MTT-containing medium was then aspirated and 200 ml DMSO was added to dissolve the formazone crystal. Optical density (OD) was measured at 570 nm in an ELISA plate reader (Fluostar Optima, BMG Labtech, Germany). Absorbance values were expressed as percentage of control.

Change in nuclear morphology of apoptotic cells
Changes in nuclear morphology of apoptotic cells were examined by fluorescence microscopy of DAPI-stained cells. In brief, 0.56106 cells were seeded in a 6-well plate and incubated (24 h) with 10 and 20 mM concentrations of PTER-ITC in presence and absence of GW9662. For this the cells were pretreated with 10 mM GW9662 for 1 h, followed by treatment with 10 and 20 mM PTER-ITC (for the next 24 h). The cells were then washed with PBS (phosphate-buffered saline) and incubated with 500 ml DAPI (0.5 mg/ml; 10 min, in the dark) and observed by fluorescence microscopy (Zeiss, Axiovert 25).  annexin V, Alexa Fluor 488 (Alexa488) and propidium iodide for cell staining in binding buffer (room temperature, 15 min in the dark). Stained cells were analyzed on a fluorescence activated cell sorter (FACS Calibur, BD Biosciences, San Jose, CA) and data were analyzed using Cell Quest 3.3 software.

Immunofluorescence staining
For immunofluorescence staining, cells were washed with PBS and fixed in 3% paraformaldehyde, permeabilized with 0.1% Triton X-100 and blocked with 1% BSA (bovine serum albumin; 30 min, room temperature). Cells were then incubated with anti-PPARc antibody (1:200 in blocking buffer; 1 h, room temperature). Finally, the cells were washed with PBS and incubated with FITC-labeled anti-rabbit secondary antibody (1:1000 in blocking buffer; 30 min, room temperature) and observed by fluorescence microscopy (Zeiss, Axiovert 25).

Luciferase assay
PPARc activity was studied by luciferase assay as described [18]. Briefly, cells were seeded at density of ,46104 cells/well in 12-well microtiter plates, and incubated overnight. Cells were then incubated in serum-free DMEM for $1 h before transfection with PPREx3-tk-Luc (three PPRE from rat acyl-CoA oxidase promoter under the control of the Herpes simplex virus thymidine kinase promoter) and Renilla-luc plasmids as an internal control. For PPAR study, cells were transfected with 25 ng pcMX-PPARa, pcMX-PPARb and pcMX-PPARc plasmids, each with 250 ng of reporter gene plasmid using Polyfect transfection reagent (Qiagen), according to instructions. Transfected cells were exposed to vehicle, various concentrations of PTER, PTER-ITC and PPAR agonist or antagonist in charcoal-stripped medium (24 h). Cells were then lysed and luciferase activity measured according to kit instructions (Promega, Madison, WI). Triplicates were measured for each experimental point; variability was ,10%. Luciferase values for each lysate were normalized to Renilla luciferase activity.

Oil Red O staining of MCF-7 cells
Approximately 105 cells were cultured on glass coverslips and treated at different PTER-ITC and rosiglitazone concentrations. After 2 days, and every 2 days thereafter, cells were switched to fresh drug-containing medium. MCF-7 cells differentiated for a total of 7 days were washed twice with PBS (pH 7.4) and fixed with 2 ml 10% formalin in PBS (30 min, room temperature). Cells were then washed twice with 2 ml distilled water and stained with 0.5% Oil Red O (Sigma, St. Louis, MO) for 10 min with gentle agitation. Excess stain was removed with 60% isopropanol and cells were washed twice with distilled water before imaging under a light microscope. Accumulated lipids were extracted in 2 ml 100% isopropanol and absorbance measured at 510 nm.

RT-PCR
Total RNA was extracted from the treated cells using an RNA isolation kit (Genei). Samples were then quantified and equal amounts of the individual treatments were transcribed with the RT-PCR kit (Genei) according to instructions. Similar treatments, followed by RNA isolation and RT-PCR were carried out three times to eliminate inter-assay variations. Primers for PPARc, PTEN and b-actin were designed using Primer 3 software and standardized in the laboratory. Primer sequences were 59-TCTGGCCCACCAACTTTGGG-39 (sense) and 59-CTTCA-CAAGCATGAACTCCA-39 (anti-sense) for PPAR-c, 59-AC-CAGG ACCAGAGGAAACCT-39 (sense) and 59-GCTAGCCTCTGGATTTGACG-39 (anti-sense) for PTEN and 59-TCACCCACACTGTGCCCCATCTACGA-39 (sense) and 59-CAGCGGA ACCGCTCATTGCCAATGG-39 (anti-sense) for b-actin. Amplification of PPARc and PTEN comprised of 29 cycles (PPARc: 94uC for 60 s, 55uC for 45 s, 72uC for 2 min; PTEN: 94uC for 60 s, 58uC for 45 s, 72uC for 2 min), and for the b-actin control: 25 cycles (94uC for 60 s, 57uC for 45 s, 72uC for 2 min). PCR conditions were optimized to maintain amplification in the linear range to avoid the plateau effect. PCR products were then separated on a 2% agarose gel and visualized in a gel documentation system (BioRad, Hercules, CA). Band intensity on gels was analyzed using ImageJ 1.43 software (NIH, Bethesda, MD) and normalized to b-actin PCR products. Each RT-PCR was carried out three times.

Western blot analysis
For western blot analysis, lysates were prepared by harvesting cells in lysis buffer [20 mM Tris pH 7.2, 5 mM EGTA, 5 mM EDTA, 0.4% (w/v) SDS and 1X protease inhibitor cocktail]. Protein was quantified with a BCA protein estimation kit (Sigma). Total protein samples (,40 mg) were analyzed on 12% polyacrylamide gels, followed by immunoblot analysis using a standard protocol. In brief, proteins were transferred to nylon membrane, which was blocked with TBS-T buffer (20 mM Tris-HCl, pH 7.5, 150 mM NaCl, 0.05% Tween-20) containing 5% skim milk powder. The blots were washed with TBS-T buffer and incubated (overnight, 4uC) in the same buffer with primary anti-PPARc, -PTEN, -survivin, -Bcl-2, -Bax caspase-9 (1:500) or -b-actin (1:1000) antibodies (all from Santa Cruz Biotechnology). Blots were then washed and incubated with HRP (horseradish peroxidase)-conjugated anti-rabbit or -mouse secondary antibody (1:20,000). Color was developed in the dark using the ECL kit (GE Healthcare, Bucks, UK) and blots were analyzed by densitometry with ImageJ 1.43 using b-actin as internal control.

Molecular docking study
Docking simulations were performed with Glide using the Maestro module of the Schrödinger suite (Suite 2011: Maestro v. 9.2, Schrödinger, New York NY). The crystal structure of PPARc bound to ligand Telmisartan was used as the starting model (PDB ID 3VN2) [43]. Using the protein preparation wizard, the complex was prepared by addition of hydrogens and sampling at neutral pH. The structure was refined with the optimized potential for liquid simulations (OPLS) 2005 force field [44] and minimized to a root mean square deviation (RMSD) of 0.30 Å . The Telmisartan binding pocket, which lies within the protein ligandbinding domain (LBD; residues 225-505), was identified on the PPARc/Telmisartan complex and the receptor grid was generated. During this process, no Van der Waal radius sampling was done; the partial charge cut-off was set at 0.25 and no constraints were enforced [45]. Ligands under study were drawn with ChemDraw [46] and 3-D structure files were generated at Online SMILES Translator and Structure File Generator (http://cactus. nci.nih.gov/services/translate/), followed by preparation with the Maestro LigPrep wizard. Each ligand was subjected to a full energy minimization in the gas phase employing OPLS2005 force field [44], with the generation of structures by different combinations of ionized states and considering all possible tautomeric states in a pH range of 5 to 9. Docking calculations were done using the Extra Precision (XP) mode of Glide [47], maintaining the receptor fixed and ligand flexible. This mode incorporates a more refined and advanced scoring function for protein-ligand docking, which gives an overall approximation of the ligand binding free energy. The function is given by where E coul is coulomb interaction energy; E vdw is Van der Waals interaction energy; E bind is binding energy and E penalty is energy due to disolvation and ligand strain. Finally, post-docking energy minimization was used to improve the geometry of the poses.

Statistical analysis
Data are expressed as mean 6 SEM and statistically evaluated with one-way ANOVA followed by the Bonferroni post hoc test using Graph Pad Prism 5.04 software (Graph Pad Software, San Diego CA). A p value of ,0.05 was considered statistically significant.

Results
PPARc is involved in PTER-ITC-induced inhibition of cell proliferation MCF-7 and MDA-MB-231 cells were treated with increasing concentrations (1-60 mM) of PTER and PTER-ITC for 24 h and cell survival was determined by MTT assay. Our data showed that treatment of these cells with PTER and PTER-ITC resulted in dose-dependent inhibition of cell proliferation, which was more pronounced after PTER-ITC treatment compared to vehicletreated control cells (Fig. 1B, C). In MCF-7 cells treated with 10 and 20 mM PTER-ITC, viable cell numbers decreased from 75% to 55%, which was about 92% and 85% respectively, after PTER treatment (Fig. 1D). Preincubation of cells with 10 mM GW9662 (a PPARc antagonist) increased cell survival from 75% to 87% in the presence of 10 mM PTER-ITC, which was 55% to 67% in the case of 20 mM PTER-ITC (p,0.05) (Fig. 1D). PTER treatment did not lead to improvement in viability when cells were pretreated with GW9662. Results were similar for MDA-MB-231 cells, in which with 10 mM GW9662 pretreatment increased cell survival from 82% to 97% in the presence of 10 mM PTER-ITC, and 70% to 87% after 20 mM PTER-ITC treatment (p, 0.05) (Fig. 1E).

Differential PPARc expression in distinct breast cancer cell lines
Three breast cancer cell lines (MCF-7, MDA-MB-231, T47D) were analyzed for PPARc expression. RT-PCR results showed that PPARc transcription was highest in MDA-MB-231 cells compared to the other two cell lines ( Fig. 2A, left). In accordance, we found that PPARc protein expression was also higher in MDA-MB-231 cells, followed by MCF-7 and T47D cell lines ( Fig. 2A, right). Based on these results, we selected MCF-7 and MD-MB-231 cells as in vitro models for the remaining part of the study.

PTER-ITC upregulates PPARc expression and activity
To examine changes in PPARc mRNA and protein expression following exposure to different drugs, we used RT-PCR, immunoblot and immunofluorescence analysis. In MCF-7 cells, the PPARc transcript level increased in response to PTER-ITC in a dose-dependent manner, which was ,1.5-fold at the highest dose tested (Fig. 2B). In contrast, PTER showed no significant increase, while the PPARc agonist rosiglitazone caused a 1.7-fold upregulation in its expression, as anticipated. Results were similar in MDA-MB-231 cells, in which PTER-ITC, PTER and rosiglitazone showed 1.6-, 1.1-and 1.8-fold increases in PPARc mRNA levels at a 20 mM concentration (Fig. 2C). This result was validated by immunoblot analysis, in which we observed a dosedependent increase in PPARc protein expression after PTER-ITC treatment in MCF-7 (2.1-to 2.8-fold) and MDA-MB-231 cells (1.5-to 2.6-fold) (Fig. 2D, E) (p,0.05). Treatment with 20 mM PTER had little or no effect, while treatment with same dose of rosiglitazone led to a significant increase in PPARc expression in MCF-7 and MDA-MB-231 cells (p,0.05). Immunofluorescence analysis of PPARc localization also showed increased nuclear accumulation of PPARc for PTER-ITC-and rosiglitazone-treated MCF-7 (Fig. 3A) and MDAMB-231 cells (Fig. 3B) compared to control cells, which was markedly inhibited by GW9662. PTER treatment led to no increase in PPARc expression or activity. These data show that PPARc expression was upregulated by PTER-ITC at both the transcriptional and translational levels.

PPARc participates in PTER-ITC-mediated upregulation of the PTEN tumor suppressor gene
To determine the effect of PTER, PTER-ITC and rosiglitazone on the expression pattern of the tumor suppressor gene PTEN, we treated MCF-7 and MDA-MB-231 cells with various concentrations of drugs for 24 h. RT-PCR and immunoblot analysis showed that PTER-ITC increased PTEN expression at both the transcriptional (Fig. 2B, C) and translational levels (Fig. 2D, E) in a dose-dependent manner (p,0.05). The most effective dose was 20 mM PTER-ITC, which caused an increase almost comparable to that of rosiglitazone. There was little or no difference in the relative level of PTEN in the PTER-treated group compared to controls (Fig. 2D, E) (p,0.05).

PTER-ITC increased PPARc and PPARb activity in MCF-7 cells
We used a luciferase reporter-based transactivation assay to study the effect of PTER-ITC on the activity of various PPAR types in breast cancer cells. Cells were transfected with plasmids encoding each PPAR protein (pcMX-PPARa, pcMX-PPARb or pcMX-PPARc) and with PPRE-tk-Luc and Renilla luciferase plasmids as internal control. Cells were then treated with PTER and PTER-ITC (24 h), followed by extraction of whole-cell lysates for analysis of luciferase activity. PTER-ITC induced PPARb and PPARc activities, but had no significant effects on PPARa ( Fig. 4A; p,0.05), whereas PTER induced PPARa activity, with no significant change in PPARb and PPARc activities ( Fig. 4A; p,0.05). We examined the specificity of PTER-ITC on PPARc and PPARb activity, using their respective agonists and antago- nists. The PPARb antagonist GSK0660 did not reverse PTER-ITC-induced PPARb activity (Fig. 4B), suggesting that the PTER-ITC effect on PPARb was non-specific. The PPARc antagonist GW9662 reversed PTER-ITC-induced PPARc activity significantly (Fig. 4C, left), as well as the activity of rosiglitazone, a PPARc agonist (Fig. 4C, right). These data suggest that PTER-ITC activity is mediated via the PPARc but not the PPARb pathway.

Effects of PTER-ITC on MCF-7 cell differentiation
PPARc activation induces cells to a more differentiated, less malignant state and causes extensive lipid accumulation in cultured breast cancer cells [30]. We thus used Oil Red O staining to test whether addition of PTER-ITC and rosiglitazone in MCF-7 cells also induces differentiation. Untreated MCF-7 cells showed nominal lipid accumulation as measured by Oil Red O staining (Fig. 4D, left). In contrast, rosiglitazone treatment (10 mM) strongly induced lipid accumulation; PTER-ITC treatment also caused a dose-dependent increase in lipid accumulation, albeit to a lesser extent than rosiglitazone (Fig. 4D). Maximum lipid accumulation was found at 5 mM PTER-ITC (Fig. 4D, right).

Molecular modeling of PPARc LBD/PTER-ITC binding
Since PTER-ITC increased PPARc transactivation by acting as a selective PPARc ligand, we used molecular docking analysis to further study PPARc LBD (ligand-binding domain)/PTER-ITC interaction at the cellular level. PTER-ITC, its parent compound (PTER), and resveratrol were docked into the PPARc LBD (see Methods); the binding mode of each ligand to PPARc LBD is shown in Fig. 5A, with their respective docking scores and interaction energies in Table 1. The terms ''XP Glidescore or docking score'' and ''Emodel'' were used to denote interactions between ligand and receptor. Based on these two scores, we observed that the PTER-ITC molecule might have better binding affinity for PPARc (Table 1). In terms of interaction with different residues, PTER-ITC showed better performance than PTER and resveratrol. In the best-docked position, PTER-ITC formed two hydrogen bonds with the receptor, involving residues His323 and Tyr327 (Table 1; Fig. 5B). In addition, through extensive hydrophobic interactions, it bound more firmly to the receptor than the other two ligands (Fig. 5C). Tyr473 is involved in hydrogen bond formation with both PTER and resveratrol, indicating a similar orientation of the two molecules, which is also evident from close analysis of their docking positions (Fig. 5A). Besides hydrogen bonds and hydrophobic interactions, PTER-ITC is also involved in the formation of p-p stacking between LBD residues His449 and Phe282 and their central benzene rings. This stacking could stabilize PTER-ITC after binding and strengthen the interaction. Similar stacking is partially observed in PTER, which involves only His449.

PPARc antagonist GW9662 inhibits PTER-ITC-induced apoptosis
We analyzed PTER-ITC apoptosis induction by flow cytometry, using annexin V and propidium iodide (PI) double staining to assess the cause of decreased cell survival after PTER-ITC  treatment. We incubated MCF-7 cells with varying concentrations of PTER-ITC, alone or with GW9662 (10 mM; 24 h). PTER-ITC treatment significantly increased the percentage of apoptotic cells, and the effect was partly attenuated by pre-incubation with GW9662 ( Fig. 6A; p,0.05). Results were similar for MDA-MB-231 cells (not shown). PTER-ITC also induced apoptosisassociated morphological changes, as cells with condensed nuclei and nuclear fragmentation were apparent after treatment (Fig. 6B), which was minimal in vehicle-treated MCF-7 and MDA-MB-231 cells. The apoptotic nuclear changes were clearly reduced in cells pre-treated with 10 mM GW9662 (Fig. 6B). These data suggest that blockade of PPARc activity blunted the druginduced cell apoptosis.

PTER-ITC induces caspase-dependent apoptosis
Apoptosis is a complex activity that mobilizes a number of molecules, and its mechanisms are classified as caspase-dependent or -independent. The caspase-dependent pathway can be further   divided into extrinsic or intrinsic pathways, determined by involvement of caspase-8 or caspase-9, respectively. Both of these pathways involve activation of caspase-3/7, which is important for inducing downstream molecules responsible for DNA cleavage. To further examine the mechanism that underlies PTER-ITCinduced death of breast cancer cells, we studied a possible role for caspase in this process by measuring the enzymatic activity of caspase-3/7, -8 and -9. We observed a gradual increase in caspase-9 and caspase-3/7 activities in MCF-7 and MDA-MB-231 cells treated with 10 and 20 mM PTER-ITC for 24 h (Fig. 7A). In contrast, there were no significant changes in caspase-8 activity in MCF-7 cells, whereas we found a dose-dependent increase in activity in MDA-MB-231 cells. Our data thus suggest that PTER-ITC induced activation of the intrinsic caspase pathway in MCF-7 cells, while it induced both extrinsic and intrinsic caspase pathways in MDA-MB-231 cells.
To determine whether caspase activation was involved in PTER-ITC-induced death of cultured breast cancer cells, we used pharmacological caspase inhibitors to test whether they protect cells from undergoing apoptosis. In the case of MDA-MB-231 cells, the general caspase inhibitor Z-VAD-FMK inhibited apoptosis most efficiently (up to 70-80%; Fig. 7B, p,0.05), suggesting that apoptosis is the predominant form of cell death induced by PTER-ITC in these cells. Z-LEHD-FMK, a specific inhibitor of caspase-9, inhibited PTER-ITC-induced apoptosis by 50-55% (p,0.05), while Z-IETD-FMK, a specific inhibitor of caspase-8, inhibited PTER-ITC-induced apoptosis by 65-70% (p,0.05). In contrast, Z-LEHD-FMK inhibited PTER-ITC-induced apoptosis by 66-70% in MCF-7 cells, while Z-IETD-FMK did not effectively block PTER-ITC-induced apoptosis in this cell line, which confirmed previous reports [41]. Our data thus demonstrate that PTER-ITC-induced apoptosis is a caspasedependent process that involves both caspase-8 and -9 in MDA-MB-231 cells and only caspase-9 in MCF-7 cells.

MAPK and JNK are involved in PTER-ITC-induced PPARc activation and apoptosis
To test for a role of MAPK (mitogen-activated protein kinase) in PTER-ITC-induced PPARc activation and apoptosis of breast cancer cells, we pre-treated MCF-7 and MDA-MB-231 cells with 20 mM ERK inhibitor (PD98059), 10 mM JNK inhibitor (SP600125) or 10 mM p38 MAPK inhibitor (SB203580) for 1 h, followed by PTER-ITC treatment for an additional 24 h. Total proteins were then isolated for analysis of PPARc expression patterns. In both breast cancer cell lines, SB203580 and SP600125 pre-treatment completely blocked PTER-ITC-induced PPARc expression, whereas pre-treatment with PD98059 or DMSO had no effect (Fig. 8A). We therefore suggest that PTER-ITC induces p38 MAPK and JNK pathways to upregulate PPARc expression in MCF-7 and MDA-MB-231 cells.
Since both p38 MAPK and JNK pathways had important roles in PTER-ITC-induced PPARc expression, we evaluated whether inhibition of either pathway protected cells from PTER-ITCinduced apoptosis. The breast cancer cells were pre-treated with 10 mM SB203580 (p38 MAPK inhibitor) or SP600125 (JNK inhibitor) for 1 h, followed by PTER-ITC treatment for an

PTER-ITC induces apoptosis by targeting PPARc-related proteins
To elucidate the mode of action of PTER-ITC as an apoptotic agent in the PPARc-dependent pathway, we studied its effect on the regulation of PPARc-related genes in both breast cancer cell lines. PTER-ITC significantly increased PPARc, PTEN and Bax, and decreased Bcl-2 expression in a dose-dependent manner both at the level of transcription (not shown) and translation (Fig. 9A,  B). Moreover, PTER-ITC significantly decreased expression of survivin, which blocks caspase-9 and -3, thereby inhibiting apoptosis.
To determine whether the increase in apoptosis and decrease in PPARc-related genes was due to PTER-ITC-induced PPARc activation, we performed two sets of experiments. First, we used the PPARc antagonist GW9662 to block PPARc pathway activation, followed by 24 h PTER-ITC treatment. Second, PPARc protein expression was knocked down in MCF-7 and MDA-MB-231 cells by transfection of PPARc siRNA, followed by 24 h PTER-ITC treatment. Our results showed that MCF-7 and MDA-MB-231 cells in both treatment protocols restored the inhibition of Bcl-2 and survivin caused by PTER-ITC alone ( Fig. 9A-D). In addition, PTER-ITC upregulated Bax and PTEN protein expression in a dose-dependent manner, which was inhibited by the PPARc antagonist or PPARc siRNA (Fig. 9A-D), indicating that PTER-ITC modulation of Bax and PTEN is PPARc-dependent. Furthermore, PTER-ITC induction of cleaved caspase-9 in both MCF-7 and MDAMB-231 cells was attenuated by GW9662 or PPARc siRNA treatment (Fig. 9). These data suggest that PTER-ITC induced PPARc expression, which subsequently enhanced expression of downstream components of this pathway, finally leading to apoptosis.

Discussion
Breast cancer is the most commonly diagnosed cancer and the second leading cause of cancer death [48]. The mortality rate of breast cancer is high because of disease recurrence, which remains the major therapeutic barrier in this cancer type. Although many cytotoxic drugs have been developed for clinical use, cancer chemotherapy is always accompanied by adverse effects, which can be fatal in some cases. Due to the lack of satisfactory treatment options for breast cancer to date, there is an urgent need to develop preventive approaches for this malignancy. There is a  growing interest in combination therapy using multiple anticancer drugs that affect several targets/pathways. A single molecule containing more than one pharmacophore, each with a different mode of action, could be beneficial for cancer treatment. Here, we studied the effectiveness of a new synthetic derivative of pterostilbene, a phytochemical isolated from Pterocarpus marsupium stem heart wood, in hormone-dependent (MCF-7) andindependent (MDA-MB-231) breast cancer cell lines.
PPARc is widely expressed in many tumors and cell lines, and has become a promising target for anticancer therapy. This nuclear receptor has a critical role in breast cancer proliferation, survival, invasion, and metastasis [13,18,20,21,[25][26][27][28]. The effectiveness of PPARc agonists as anticancer agents has been examined in various cancers including colon, breast, lung, ovary and prostate [49]. We tested whether PTER-ITC mediates its anti-proliferative and pro-apoptotic effects in breast cancer cells through activation of the PPARc signaling cascade. Our results showed that PTER-ITC activated PPARc expression in a dosedependent manner, followed by downregulation of its antiapoptotic genes (Bcl-2 and survivin) to induce noteworthy levels of apoptosis in hormone-dependent (MCF-7) and -independent (MDA-MB-231) breast cancer cells.
The PTER-ITC conjugate can be considered more advantageous than existing PPARc ligands such as rosiglitazone or pioglitazone for breast cancer treatment, as PTER-ITC causes more pronounced cell death at a much lower dose than other ligands [50][51][52]. In addition, most (if not all) the other ligands are estrogenic in nature [53], and could thus act as positive factors for ER-dependent breast, ovary and uterine cancers, whereas PTER-ITC is anti-estrogenic at the dose used for this study. Considering these two major points, we consider that the drug could be used at much lower concentrations, which might help reduce the side effects reported for most other PPARc ligands. PTER-ITC molecule nonetheless requires further validation before use in clinical trials that target the PPARc pathway.
The most important characteristic of a cancer cell is its ability to sustain proliferation [54]. The pathways that control proliferation in normal cells are altered in most cancers [55]. We thus analyzed the PTER-ITC effect on proliferation of breast cancer cells, and found that PTER-ITC caused significant, dose-dependent inhibition of breast cancer cell growth in vitro. This effect was partially reversed, however, when PTER-ITC was combined with PPARc antagonists. This result suggests that the PTER-ITC anticancer effects are mediated through the PPARc activation pathway. These data coincide with findings in several in vivo and in vitro studies in which PPARc agonists such as rosiglitazone or troglitazone decreased proliferation of breast cancer cell lines, mediated in part by a PPARc-dependent mechanism [26,56].
To elucidate the molecular mechanisms that underlie the anticancer effects observed for PTER-ITC, we studied its effect on activation of PPARc. To the best of our knowledge, this is the first report showing PTER-ITC participation in the PPARc-dependent signaling pathway. Our data show that PTER-ITC increased PPARc transcriptional and translational activity in MCF-7 and MDA-MB-231 cells. To establish the essential role of PTER-ITC in PPARc-mediated apoptosis of breast cancer cells, we used PPARc siRNA and its drug antagonist to inhibit PPARc signaling, and demonstrated apoptosis prevention and caspase activation. We also observed an increase in PPARb activity after PTER-ITC treatment, with no significant reduction after antagonist treatment, suggesting that the increase was non-specific. Although some earlier studies reported involvement of PPARb activity in tumorigenesis, many others contradicted this idea. The PPARb ligand GW501516 was reported to promote human hepatocellular growth [57], although another study showed that certain PPARb ligands such as GW0742 and GW501516 reduced growth of MCF-7 and UACC903 cell lines [58]. The role of PPARb in cancer therapeutics is therefore complex and not yet fully defined [59]. Hence the relationship between PTER-ITC and PPARb could provide an alternative platform to study the involvement of this pathway in cancer therapy.
PPARc is a phosphoprotein, and many kinase pathways, such as cAMP-dependent protein kinase (PKA), AMP-activated protein kinase (AMPK) and mitogen-activated protein kinase (MAPK) such as ERK, p38 and JNK, have been implicated in the regulation of its phosphorylation [60,61]. Phosphorylation notably inhibits PPARc ligand-independent and -dependent transcriptional activation [60,61]. Research showed that PPARc agonists activate different MAPK subfamilies, depending on cell type [62][63][64][65] and that these kinases are involved in cell death [66][67][68][69]. The role of MAPK signaling pathways in cell death induced by PPARc agonists is controversial. According to certain studies, PPARc agonist-induced ERK activation mediates anti-apoptotic signaling [64], while others showed its involvement in inducing cell death [66,70]. p38 activation by PPARc agonists is also reported to be regulated differently in various cell types. PPARc agonists induce p38 activation, leading to apoptosis of cancer cells have been reported in chondrocytes [64], human lung cells [68], liver epithelial cells [62] and skeletal muscle [71]. This coincides with our data, where using pharmaceutical inhibitors, we show that activation of p38 and JNK pathways, but not of ERK, is necessary and sufficient to phosphorylate PPARc and cause subsequent apoptosis in the breast cancer cell lines studied. At present, we do not know whether PTER-ITC activates p38 and JNK directly, or if it activates other cellular kinase pathways such as PKA and AMPK, which in turn could activate MAPK. Further validation is needed to conclusively establish the pathway(s) involved.
PTEN is a tumor suppressor gene involved in the regulation of cell survival signaling through the phosphatidylinositol 3-kinase (PI3K)/Akt pathway [72]. PI3K/Akt signaling is required for an extremely diverse array of cellular activities that participate mainly in growth, proliferation, apoptosis and survival mechanisms [73,74]. Activated Akt protects cells from apoptotic death by inactivating compounds of the cell death machinery such as procaspases [73]. PTEN exercises its role as a tumor suppressor by antagonizing the PI3K/Akt pathway [73]. The PPARc-dependent increase in PTEN caused by PTER-ITC in our experiments not only indicates that the tumor suppressor gene contributes to the growth-inhibitory activities of the compound, but might also trigger its pro-apoptotic actions.
Our results further showed that PTER-ITC downregulated PPARc-related genes, including Bcl-2 and survivin. These genes are commonly associated with increased resistance to apoptosis in human cancer cells [75]. PTER-ITC-induced PPARc activation was reduced in the presence of GW9662, together with reversal of decreased survivin and Bcl-2 levels. Furthermore, molecular docking analysis suggested that PTER-ITC could interact with amino acid residues within the PPARc-binding domain, including five polar and eight non-polar residues within the PPARc ligandbinding pocket that are reported to be critical for its activity. Together these results suggest that PTER-ITC can be considered a PPARc agonist, and the survivin and Bcl-2 decrease is due to activation of the PPARc pathway by PTER-ITC.
Two cellular pathways, differentiation and apoptosis, are the main focus in the development of anti-cancer therapies. Induction of differentiation is one potent mechanism by which some cancer therapeutic and chemopreventive agents act [76][77][78]. Lipid accumulation in MCF-7 cells is supported by the fact that tamoxifen and a few other anti-cancer agents such as ansamycins and suberoylanilide hydroxamic acid induce high lipid production (as high as 5-fold in the case of ansamycins) and by triglyceride accumulation, which results in MCF-7 cell differentiation to a more epithelial-like morphology [79][80][81]. In a previous study, we showed that long-term exposure to PTER causes growth arrest in MCF-7 cells, which might be linked to mammary carcinoma cell differentiation into normal epithelial cell-like morphology and activation of autophagy [38]. In the present study, PTER-ITC also caused differentiation of MCF-7 cells, albeit to a higher level compared to its parent compound PTER than previously reported [38]. Based on these data, it can thus be suggested that PTER-ITC inhibits MCF-7 cell growth mainly through apoptosis, while it can also induce differentiation of these breast cancer cells.

Conclusions
In conclusion, this study highlights the anticancer effects of the novel conjugate of PTER and ITC, and shows that the mechanism involves activation of the PPARc pathway via PTER-ITC binding to the receptor, which affects its regulated gene products (Fig. 10). PTER-ITC induces apoptosis by enhancing expression of PPARc genes at both transcriptional and translational levels, which appears to be triggered at least in part by modulation of PTEN. In addition, activation of caspase-9 and downregulation of Bcl-2 and survivin contribute to PTER-ITC-induced cell death. PTER-ITC exhibits differentiation-promoting as well as anti-proliferative effects on MCF-7 cells. Together these results suggest that the PTER-ITC conjugate acts as a PPARc agonist and is a promising candidate for cancer therapy, alone or in combination with existing therapies. These preliminary data show that further studies are warranted in in vitro and in vivo models to elucidate the exact mode of action responsible for the effects of this compound.