Our previous studies revealed that MAZ is a critical transcriptional regulator of PPARγ1 in MCF-7 breast cancer cells 37. We also showed that both PPARγ1 and MAZ are highly expressed in MCF-7 cells as compared to normal mammary epithelial cells (HMEC) and tumor-specific expression of PPARγ1 is MAZ dependent 37. To evaluate expression of PPARγ1 within other breast cancer cell lines, whole cell lysates from a panel of eight different breast cancer cell lines originated from heterogeneous tumors (ranging from adenocarcinoma to metastatic ductal carcinoma) were examined by Western blot analysis. HMEC cells were used as a control. Figure 1A shows the representative immunoblot for PPARγ1. To demonstrate the reproducibility of these data, statistical analysis of three Western blots was performed (Fig. 1B). The results showed that PPARγ1 is expressed at significantly higher level in cancer cell lines as compared to HMEC.

As discussed above, evaluation of PPARγ1 as a potential breast cancer therapy target revealed the complexity of PPARγ1 signaling in cancer. The mechanism of PPARγ1 activation during cancer development and the functional role of this event in tumorigenesis still remain unclear. To address these questions and elucidate the role of PPARγ1 activation in cancer, we utilized the advantages of shRNA techniques. A set of five shRNAs for each of PPARγ or MAZ gene was purchased from The RNAi Consortium (TRC). Each shRNA was evaluated using Western blot analysis (data is not shown). The most efficient shRNA for each gene was chosen for further investigation. MCF-7 cells were transiently transfected with PPARγ and MAZ shRNAs, as well as with a scrambled shRNA as control. MCF-7 cells treated only with the transfection reagent were also used as a control. To evaluate the specificity of shRNAs to their target genes and the extent of PPARγ1 down-regulation when either PPARγ or MAZ shRNA were applied to MCF-7 cells, Real-time PCR, Western blot analysis and Luciferase assay were performed. Real-time PCR data revealed that both PPARγ and MAZ shRNAs are highly specific to their targets and their application to the cells leads to a significant decrease in PPARγ1 or MAZ mRNA levels respectively (Fig. 2A). We also tested whether the observed changes in PPARγ1 or MAZ mRNA lead to changes in PPARγ1 protein expression. Figure 2B shows the representative immunoblot for PPARγ1 and demonstrates the level of PPARγ1 down-regulation by both PPARγ and MAZ shRNAs as compared to controls. The statistical evaluation of three different Western blots showed that the direct inhibition of PPARγ1 by PPARγ shRNA resulted in an average 50 percent decrease in PPARγ1 expression. A lower level of inhibition was observed when PPARγ1 knocked-down was achieved via down-regulation of MAZ expression. This was anticipated since we believe transcription factors other than MAZ are involved in regulation of PPARγ1 in cancer cells.

It is known that PPARγ activates gene transcription by interacting with a Peroxisome-Proliferator Response Element (PPRE) located within the promoter sequence of target genes 2. To confirm that shRNAs-mediated down-regulation not only affects expression but also activity of PPARγ1, a PPRE functional response was measured. MCF-7 cells were transfected with a 3XPPRE-mTK-pGL3 reporter plasmid and then co-transfected with scrambled, PPARγ, or MAZ shRNA expression plasmids. Following transfection, cells were treated with 10 μM Rosiglitazone (Rosi), a well known PPARγ agonist 39. Cells were lysed 24 hours after the second transfection and a Luciferase assay was performed. Data demonstrated that direct down-regulation of PPARγ1 expression by PPARγ shRNA led to a significant decrease in PPRE-mediated reporter activity in both Rosi treated and untreated MCF-7 cells (Fig. 2C). This confirms that PPARγ shRNA is specific and efficient for the inhibition of PPARγ1 expression and activity. Moreover, the fact that PPRE activity falls below the control level when PPARγ shRNA is applied to the cells is additional evidence for endogenous transactivation of PPARγ1 in breast cancer cells. Although we observed a decrease in reporter activation when MAZ shRNA was transfected to the cells, these changes were not statistically significant, suggesting the complexity of PPARγ transcriptional regulation and that the knock-down of MAZ seen in transient transfection assays is not sufficient to block PPRE-mediated reporter activity.

Since one of the most important characteristics of tumor development is enhanced cell growth, we tested whether inhibition of PPARγ1 expression affects cellular proliferation in breast cancer cells. MCF-7 cells were transiently transfected with scrambled, PPARγ, or MAZ shRNA and the rate of BrdU incorporation during DNA synthesis was assessed by using the BrdU proliferation assay (Roche). The results revealed that down-regulation of PPARγ1 by both PPARγ and MAZ shRNAs significantly decreased cellular proliferation in MCF-7 breast cancer cells (Fig. 3A). To confirm these results, we used a different approach to inhibit endogenous activity of PPARγ1. MCF-7 cells were transiently transfected with a vector driving the expression of a dominant-negative mutant of PPARγ, Δ462, or an empty vector as control, and then a BrdU proliferation assay was performed. Inhibition of endogenous PPARγ1 activity using the Δ462 mutant caused a decrease in cellular proliferation in MCF-7 cancer cells (Fig. 3B). To test whether endogenous activation of PPARγ1 plays a similar role in other types of breast cancer cells, the same experiment was performed using the breast cancer cell line, T47D. The level of PPARγ1 expression in this cell line was evaluated using Western blot analysis (Fig. 3D). T47D cancer cells were transiently transfected with either a control vector or a Δ462 expression vector and then a BrdU proliferation assay was performed. Inhibition of PPARγ1 activity in these cells also led to a significant decrease in cellular proliferation (Fig. 3C). To confirm that the observed changes in cellular proliferation in both MCF-7 and T47D cell lines are indeed in response to the inhibition of PPARγ1 activity in a dominant-negative manner by Δ462, a Luciferase assay was performed. Cells were transfected with 3XPPRE-mTK-pGL3 reporter plasmid and then co-transfected with control or Δ462 expression plasmids. Following transfection, cells were treated with 10 μM Rosi. In Rosi treated and untreated cells, application of a Δ462 mutant resulted in a significantly lower level of PPRE-mediated reporter activity (Fig. 3E), thus, demonstrating that Δ462 efficiently inhibits endogenous activity of PPARγ1 in MCF-7 and T47D cancer cells.

To elucidate the mechanism by which down-regulation of PPARγ1 expression leads to inhibition of cellular proliferation in MCF-7 cells, fluorescence-activated cell sorting (FACS) was performed. Analysis of cell cycle distribution (Fig. 4) revealed that PPARγ1 down-regulation by PPARγ or MAZ shRNA primarily affects cell transition from G₁ to S-phase in MCF-7 cells. Approximately 25 percent fewer cells entered the S-phase when PPARγ1 expression was suppressed directly or indirectly. This is consistent with data from the BrdU proliferation assay. Therefore, these results confirm our hypothesis and demonstrate that an increase in PPARγ1 expression and its endogenous transactivation play an important functional role in promoting cellular proliferation in breast cancer cells.

FACS analysis allowed us to assess changes in apoptosis as well. Interestingly, the level of apoptosis (Fig. 4D) and the percentage of debris and aggregates (data not shown) in cells transfected with either PPARγ or MAZ shRNA was significantly higher than in control cells or scrambled shRNA transfected cells. This observation suggests that in addition to its involvement in regulation of proliferation, PPARγ1 may also be involved in regulation of apoptosis in MCF-7 cells.

To evaluate data from FACS analysis (Fig. 4D) and determine whether changes in PPARγ1 expression can affect apoptosis, a Cell Death Detection ELISA assay was performed which distinguishes between necrotic and apoptotic cell death. MCF-7 cells were transfected with scrambled, PPARγ, or MAZ shRNA using the same transfection protocol and time points as for the proliferation assay, FACS, and protein analysis. The results showed no significant difference in necrotic cell death between PPARγ knock-down and control cells (data not shown). However, inhibition of PPARγ1 expression in MCF-7 cells using PPARγ or MAZ shRNA resulted in a significant increase in apoptosis as compared to control (Fig. 5A). Interestingly, changes caused by down-regulation of PPARγ1 directly by using PPARγ shRNA were more dramatic than through knock-down of MAZ. This suggests that MAZ probably contributes to regulation of apoptosis mostly through its mediation of PPARγ1 signaling in cancer cells. To confirm that down-regulation of PPARγ1 expression induces apoptosis, the expression of Bcl2, an anti-apoptotic protein, which has been shown to promote the survival of cancer cells 40 was evaluated. Statistical analysis of densitometry for four Western blots demonstrated that Bcl2 expression was significantly higher in control and MCF-7 cells transfected with scrambled shRNA versus cells transfected with PPARγ or MAZ shRNA (Fig. 5B). We also assessed changes in poly (ADP-ribose) polymerase-1, PARP-1, cleavage in cells transfected with PPARγ or MAZ shRNA as compared to controls. PARP-1 cleavage by caspases is a well-known marker for apoptosis 41. Statistical evaluation of densitometry for three Western blots (Fig. 5C) showed a significant increase in an 89 kDa C-terminal fragment, the product of PARP-1 proteolysis, when PPARγ shRNA was applied to MCF-7 cells. This data suggests that an increase in PPARγ1 expression followed by transactivation during cancer development may be an important factor that contributes not only to acceleration of cellular proliferation but also to cell evasion from apoptosis.

MCF-7 and T47D breast cancer cells were obtained from the American Type Culture Collection (Rockville, MD). Cells were cultured in modified DMEM (Gibco BRL, Gaithersburg, MD) supplemented with 10% fetal bovine serum (Hyclone). Normal human mammary epithelial cells (HMEC) (Cambrex) were cultured in MEGM^® with SingleQuot^® supplements. All cell lines were grown in media lacking phenol red at 37°C in a 5% CO₂ atmosphere.

The whole cell lysates were prepared using passive lysis buffer. Concentrations were determined using a Bradford Assay (BioRad). 30 μg of total cell lysate per sample was run on a 10% or 12% SDS polyacrylamide gel. The proteins were transferred to a nitrocellulose membrane, blocked in 5% TBST, and incubated at 4°C overnight with primary antibody. The membrane was then washed and incubated for 4 hours with secondary IgG-HRP antibody. After incubation the membrane was washed, incubated with Chemiluminescence substrate (Pierce) for 5 min, and expose to film. The following primary antibodies were used: PPARγ mouse monoclonal IgG antibody 1:200 dilution (Santa Cruz Biotechnology, sc-7273), Bcl2 mouse monoclonal antibody 1:1000 dilution (Santa Cruz Biotechnology, sc-509), PARP-1 rabbit polyclonal antibody 1:1000 dilution (Santa Cruz Biotechnology, sc-2578). Appropriate secondary goat anti-mouse (Santa Cruz Biotechnology, sc-2055), or bovine anti-goat (Santa Cruz Biotechnology, sc-2378), or goat anti-rabbit secondary antibody 1:1000 dilution (Santa Cruz Biotechnology, sc-2004) antibodies 1:1000 dilution were applied. Anti-actin raised in rabbit 1:2000 dilution (Sigma, A 5060) and goat anti-rabbit secondary antibody 1:1000 dilution (Santa Cruz Biotechnology, sc-2004) were used to visualize actin. Western Blot Stripping Buffer (Pierce, # 21059) was used to restore membranes.

The set of five shRNAs for PPARγ1 (TRCN 0000001670-74) and MAZ (TRCN 0000015343-47) genes as well as scrambled shRNA and non-hairpin TRC controls were purchased from The RNAi consortium (TRC) Human shRNA Library (Open Biosystems). The shRNA construct includes a hairpin of 21 base pairs, a sense and antisense stem, and a 6 base-pair loop. Each hairpin sequence is cloned into a lentoviral vector (pLKo1). Based on structural evaluation and Western blot analysis the most efficient shRNAs for PPARγ1 (TRCN 0000001672) and MAZ (TRCN 0000015345) were chosen for transient transfections.

The dominant-negative PPARγ1 mutant was a kind gift of Dr. Stephen O'Rahilly and Dr. V. Krishna K Chatterjee, Department of Medicine, Addenbrook's Hospital, Cambridge, U.K. Sequence analysis revealed a single base deletion introducing a premature stop codon (5'-₁₃₈₀ GACAGACTGA₁₃₉₀-3') leading to translation of protein truncated just before the AF-2 domain.

Cells were transiently transfected with 3.6 μg of pGL3 plasmid containing 3XPPRE-mTK-Luc and Renilla (Allred, 2005) per 24-well plate and then co-transfected with scrambled, PPARγ or MAZ shRNAs, Δ462, or control plasmids using FuGENE transfection reagent (Roche). 4 hours after transfection cells were subsequently treated with 10 μM Rosi for 20 hours. Cells were lysed in 80 μl of passive lysis buffer and treated according to manufacture's instructions (Promega dual luciferase assay kit). Luminometry was performed on a Berthold Technologies Lumat 9507 (Wildbad, Germany). Results were calculated as raw Luciferase units divided by raw Renilla units (RLU's). Data is presented as mean fold changes in treated cells as compared to control cells.

Total mRNA was isolated using an RNeasy Mini Kit (Qiagen, CA) according to manufactures instructions. Real-time PCR was performed on total RNA using the TaqMan One-Step RT-PCR Master Mix Reagents Kit (Applied Biosystems). The pre-optimized primers and probes for MAZ, PPARγ1, and 18S were purchased from Applied Biosystems.

MCF-7 cells were seeded at 0.1 × 10⁴ cells/well in 96-well tissue culture plates. Cells were transiently transfected on the second and third day using 0.05 μg of plasmid and 0.3 μl of FuGENE 6 transfection reagent (Roche) per well. Control MCF-7 cells were treated with FuGENE6 only. 16 wells per each shRNA and control were used. The same experimental set-up was used when cells were transfected with a dominant-negative form of PPARγ1, Δ462. The media was changed before the second transfection. Cell Proliferation ELISA, BrdU (colometric) (Roche) was performed on the fifth day according to the manufacture instructions.

The same protocol as for the proliferation assay was used to plate and transfect MCF-7 cells. A Cell Death Detection ELISA (Roche) was performed on the fifth day. The assay is based on quantitative sandwich-enzyme-immunoassay-principle and uses mouse monoclonal antibodies directed against DNA and histones. Cells were lysed in a 96-well plate, centrifuged, and 20 μl of supernatant was transferred into streptavidin-coated wells. A mixture of antibodies was added and the plate was then incubated for 2 hours. The unbound components were removed by washing. ABTS substrate was added and the amount of mono- and oligonucleosomes were measured photometrically using the ELISA-plate reader according manufacture instructions (Roche).

The DNA content of control and shRNAs transfected MCF-7 cells was analyzed using a detergent-trypsin method (Vindelov, 1983). MCF-7 cells were seeded at 1 × 10⁶ cells in 100 mm culture plates. Cells were transiently transfected on the second and third day with 6 μg of plasmid using 18 μl of FuGENE6 transfection reagent (Roche). On the fifth day the propidium iodide labeling procedure and fluorescence-activated cell sorting (FACS) using Mod FitLT V.3.1 software was performed (University of Kentucky Flow Cytometry Facility).

Data was analyzed by a two-way analysis of variance (ANOVA) using the StatServer 6.1(Insightful, Seattle, WA) from the server maintained by the University of Kentucky's Department of Statistics. In every 2-way ANOVA, Tukey's pair-wise comparison test was used post-hoc. P-values of less than 0.05 were considered to be significant. One-way ANOVA with Fisher's LSD or Tukey's pair-wise comparison post-hoc test were also used where appropriate. When appropriate, Student's t-test was also used for data analysis on Microsoft Excel.