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Peroxisome proliferator-activated receptor β expression in human breast epithelial cell lines of tumorigenic and non-tumorigenic origin
The International Journal of Biochemistry & Cell Biology 34 (2002) 1051–1058
Peroxisome proliferator-activated receptor  expression in human breast epithelial cell lines of tumorigenic and non-tumorigenic origin Kate M. Suchanek, Fiona J. May, Won Jae Lee, Nicola A. Holman, Sarah J. Roberts-Thomson∗ School of Pharmacy, The University of Queensland, St. Lucia, QLD 4072, Australia Received 23 November 2001; received in revised form 6 February 2002; accepted 7 February 2002
Abstract Peroxisome proliferator-activated receptor  (PPAR) is a member of the nuclear hormone receptor superfamily and is a ligand activated transcription factor, although the precise genes that it regulates and its physiological and pathophysiological role remain unclear. In view of the association of PPAR with colon cancer and increased mRNA levels of PPAR in colon tumours we sought in this study to examine the expression of PPAR in human breast epithelial cells of tumorigenic (MCF-7 and MDA-MB-231) and non-tumorigenic origin (MCF-10A). Using quantitative RT-PCR we measured PPAR mRNA levels in MCF-7, MDA-MB-231 and MCF-10A cells at various stages in culture. After serum-deprivation, MDA-MB-231 and MCF-10A cells had a 4.2- and 3.8-fold statistically greater expression of PPAR compared with MCF-7 cells. The tumorigenic cell lines also exhibited a significantly greater level of PPAR mRNA after serum deprivation compared with subconfluence whereas such an effect was not observed in non-tumorigenic MCF-10A cells. The expression of PPAR was inducible upon exposure to the PPAR ligand bezafibrate. Our results suggest that unlike colon cancer, PPAR overexpression is not an inherent property of breast cancer cell lines. However, the dynamic changes in PPAR mRNA expression and the ability of PPAR in the MCF-7 cells to respond to ligand indicates that PPAR may play a role in mammary gland carcinogenesis through activation of downstream genes via endogenous fatty acid ligands or exogenous agonists. © 2002 Elsevier Science Ltd. All rights reserved. Keywords: PPAR; Breast; Human; Cell lines
1. Introduction Abbreviations: ATCC, American type culture collection; CT , threshold cycle; DMEM, Dulbecco’s modified eagle medium; FBS, fetal bovine serum; Real time RT-PCR, real time reverse transcriptase-PCR; PPAR, peroxisome proliferator-activated receptor; RTPCR, reverse transcriptase-PCR; S.E.M., standard error of the mean ∗ Corresponding author. Tel.: +61-7-3365-3193; fax: +61-7-3365-1688. E-mail address: [email protected] (S.J. Roberts-Thomson).
Peroxisome proliferator-activated receptor  (PPAR; NR1C2) is a member of the nuclear hormone receptor superfamily that includes the thyroid hormone, Vitamin D and retinoic acid receptors [1]. Members of the PPAR family also include PPAR␣, PPAR␥1, PPAR␥2 and PPAR␥3 [2–4]. While PPAR␣ and PPAR␥ have clearly defined roles
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in controlling lipid and glucose homeostasis (reviewed in [5]) the physiological role of PPAR remains less defined with elucidation of its function hindered by a lack of a widely available specific agonist. PPAR is expressed in humans at similar levels in all tissues so far examined [6]. Its expression in rodent tissues is also ubiquitous although its expression level is often greater than that of PPAR␣ or ␥ [7]. Endogenous ligands of PPAR include the polyunsaturated fatty acids arachidonic acid and linoleic acid [8], which also act as agonists for PPAR␣ and ␥ [8–10]. PPAR has been associated with a number of biological functions including embryo implantation [11], osteoclastic bone resorption [12], lipid homeostasis [13–15], skin wound healing [16] and colon cancer [17–19]. Using PPAR-null mice in vivo evidence has been presented linking PPAR with roles in development, myelination of the corpus callosum, lipid metabolism and epidermal cell proliferation [20]. Significantly, studies using the PPAR-null mice [20] highlight a role for PPAR in cell cycle control, a finding that supports an earlier study suggesting a link between PPAR and human colon cancer [19]. In four individual patients with colorectal cancer, increased PPAR mRNA expression was seen in cancer tissue compared with normal colon epithelium, suggesting that the level of PPAR expression may be a factor influencing colon carcinogenesis [19]. Indeed, a total absence of PPAR in a human colorectal cancer cell line reduced the ability of the cells to form tumours in nude mice [18]. PPAR is expressed in the mammary gland of the mouse [21] as well as cells of epithelial origin [22]. Despite the ubiquitous nature of PPAR expression and its association with tumorigenesis, there are no studies examining the expression of PPAR in human breast epithelial cells. We have sought in this study to examine the expression of PPAR in two human breast cancer epithelial cell lines and a non-tumorigenic breast epithelial cell line and assess changes in PPAR expression levels during proliferation and in response to agonist stimulation as a first step in characterising any potential role for this receptor in mammary gland tumorigenesis.
2. Materials and methods 2.1. Materials Cell culture media, antibiotics and fetal bovine serum (FBS) were obtained from Life Technologies (Mulgrave, Australia). MCF-7, MDA-MB-231 and MCF-10A cell lines were purchased from the American type culture collection (ATCC). Six well plates were purchased from Nalge Nunc International (IL, USA) and bezafibrate was purchased from Sigma (Castle Hill, Australia). The human brain cDNA library was the kind gift of Professor M.E. McManus (The University of Queensland, Australia). All other reagents and materials were purchased from Sigma. 2.2. Cell culture Cells were cultured in Dulbecco’s modified eagle medium (DMEM) supplemented with 10% FBS, glutamine (2 mM), penicillin G (100 units/ml) and streptomycin sulphate (100 mg/ml) in 6 well plates and maintained at 37 ◦ C and 5% CO2 . Prior to confluence, cells were trypsinised with a 0.05% trypsin/0.53 mM EDTA solution and resuspended in fresh growth media before plating, at a ratio of 1:6, onto a new growth surface. RNA was isolated from subconfluent proliferating cells and confluent cells (>95% confluence). For studies involving FBS deprivation, confluent cells were cultured for 72 h in FBS-free media prior to RNA isolation. For studies involving the addition of bezafibrate (50 M) MCF-7 cells were allowed to reach confluence in normal growth media then cultured in the absence of FBS for 48 h prior to the addition of bezafibrate (in DMSO) in fresh FBS-free media for 24 h. Control cells were incubated in the presence of DMSO 0.12%. All experiments were performed in triplicate. 2.3. RNA isolation The QIAGEN RNeasyTM mini kit (QIAGEN, Clifton Hill, Vic., Australia) was used to isolate total RNA and included an on-column DNase treatment (QIAGEN). RNA was resuspended in RNase free water and stored at −80 ◦ C until required. RNA was quantitated by absorbance at 260 nm while purity was determined by the ratio of absorbance at 260 and 280 nm.
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2.4. Reverse transcriptase-PCR The access RT-PCR system (Promega, Sydney, Australia) was used for reverse transcriptase-PCR (RT-PCR). Following reverse transcription at 48 ◦ C for 45 min using gene-specific primers a Stepdown PCR protocol was followed [23]. The annealing temperature ranged from 65 to 49 ◦ C for 30 s with extension at 68 ◦ C for 30 s. Each reaction contained 0.5 g RNA, Mg2+ (1.5 mM for MDA-MB-231 and brain cDNA library; 2 mM for MCF-7 and -10A) primers 0.5 M and dNTPs 0.2 mM. Primers for human PPAR were as follows; forward: 5 -AAAGAAGGCCCGCAGCAT3 , reverse: 5-CCTTCTCTGCCTGCCACAAT-3 and were designed to amplify a 92 bp product. For all reactions a human brain cDNA library was used as a positive control and reactions run in the absence of the reverse transcriptase enzyme were used as negative controls. RT-PCR reactions were analysed by agarose gel electrophoresis and bands were excised from the gel and sequenced to confirm identity. 2.5. Real time RT-PCR Primers and probes for real time RT-PCR were designed using the software Primer Express Version 1.0 (Applied Biosystems, Foster City, USA). Primers for PPAR were as described above. The probe (5 -TGCTGCGCTGCACCCGCT-3 ) had the fluorescent dye FAM (6-carboxy-fluorescin) attached to the 5 end and the quencher dye TAMRA (6-carboxy-tetramethylrhodamine) at the 3 end. Probe and primers for the standard 18S rRNA were purchased from Applied Biosystems. RNA isolated from MCF-7 cells cultured in normal growth media was used to obtain the standard curve using 0.1–100 ng total RNA for PPAR amplification and 0.001–1 ng total RNA for 18S rRNA amplification. Equal amplification efficiency of the target and standard amplicons was determined by linear regression analysis and examination of R2 using Microsoft® Excel (Microsoft Corporation, Washington, USA). Using the Taqman® Gold RT-PCR kit (Applied Biosystems), each real time RT-PCR reaction (25 ml) contained 1 × Taqman A buffer, 5.5 mM magnesium chloride, 300 mM dATP, dCTP and dGTP, 600 mM dUTP, AmpliTaq Gold DNA polymerase 0.625 units, MultiScribe reverse transcriptase 0.625 units, forward
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and reverse primers (PPAR 900 nM, 18S rRNA 50 nM), RNA 25 ng (PPAR) or 0.25 ng (18S rRNA), probe (PPAR 200 nM, 18S rRNA 50 nM) and RNase inhibitor 10 units. RNA was replaced with RNase free water in no template controls. Reactions were cycled in an ABI PRISM 7700 Sequence Detector (Applied Biosystems) using the following conditions; 48 ◦ C for 30 min, 95 ◦ C for 10 min then 45 cycles of 95 ◦ C for 15 s and 60 ◦ C for 1 min, followed by 25 ◦ C for 10 min. For real time RT-PCR CT values (cycle number at which amplicon product is first detected above baseline) were used to calculate CT for each sample (CT PPAR␣ − C T 18S rRNA). To allow comparison between cell lines, a CT value was calculated by normalising values to MDA-MB-231, 72 h FBS deprivation. The MDA-MB-231 cell line was chosen for normalisation due to its greater levels of PPAR mRNA compared with MCF-10A and -7 cells cultured under the same conditions. The 72 h time point was chosen, as this time point was used to compare cell lines. Fold change is calculated from CT using the formula 2−CT [24]. Statistical analysis of samples was performed using SigmaStat® (SPSS Science, Chicago, IL, USA). Results were analysed using oneway ANOVA with the Tukey Test for all pairwise multiple comparison for normally distributed data of equal variance, or where data were not normally distributed the Kruskal–Wallis ANOVA on Ranks with Dunn’s Method of Multiple Comparison was used. Significant (P < 0.05) values are indicated in the result section.
3. Results 3.1. Expression of PPARβ in breast epithelial cell lines Using RT-PCR we showed that PPAR is expressed in human breast epithelial cell lines (Fig. 1): tumorigenic MCF-7 and MDA-MB-231 [25] and non-tumorigenic MCF-10A [26]. 3.2. Development of a quantitative assay for PPARβ mRNA in breast epithelial cell lines A real time RT-PCR assay was developed to quantify the mRNA level of PPAR in relation to 18S rRNA
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Fig. 1. RT-PCR amplification of a 92 bp fragment of PPAR in breast epithelial cell lines. The PPAR fragment was amplified as outlined in Section 2: lane 1, marker; lane 2, human brain cDNA control; lane 3, MDA-MB-231; lane 4, MDA-MB-231, -reverse transcriptase (RT); lane 5, MCF-7; lane 6, MCF-7, -RT; lane 7, MCF-10A; lane 8, MCF-10A, -RT.
levels. We initially validated the real time RT-PCR assay to ensure that our target and internal standard amplicons were amplified with the same efficiency. Relative comparisons can be made with regard to amplicon levels when efficiencies are equal [27]. To determine this we examined the relationship between log (total RNA) and CT values (CT values are defined as the fraction cycle number at which fluorescence passes
a predetermined threshold (set at 10 times the standard deviation of the baseline [28]) for the amplification of PPAR and 18S rRNA (Fig. 2). If the slope of a plot of log (input amount) versus CT (CT PPAR − C T 18S rRNA) has an absolute value of <0.1 this suggests an approximate equal amplification efficiency. We determined that equal amplification efficiency lies between 1 and 100 ng total RNA for PPAR and
Fig. 2. Standard curve for the real time RT-PCR amplification of PPAR and 18S rRNA in MCF-7 human breast cancer epithelial cells. Each data point represents the mean ± S.D. (n = 3). The x-axis shows the concentration of total RNA on a logarithmic scale. The RNA was diluted 100-fold for the measurement of 18S rRNA.
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Fig. 3. PPAR mRNA expression in MDA-MB-231, MCF-7 and -10A human breast epithelial cell lines by real time RT-PCR with reference to 18S rRNA. Expression levels were quantitated in subconfluent proliferating cells and at >95% confluence (confluent cells) and after 72 h in FBS deprivation as described in Section 2. CT have been normalised to MDA-MB-231, 72 h FBS deprivation to give a CT value for each group. The MDA-MB-231 cell line was chosen for normalisation due to its greater levels of PPAR mRNA compared with MCF-10A and -7 cells cultured under the same conditions. Each bar represents mean ± S.E.M. (n: subconfluence, confluence, 72 h FBS deprivation; MDA-MB-231 and MCF-10A; 12, 21, 33; MCF-7, 12, 11, 22, respectively). Symbol (∗) denotes significant difference. After 72 h serum deprivation, MDA-MB-231 and MCF-10A cells had 4.2- and 3.8-fold higher levels of PPAR mRNA, respectively, than MCF-7 cells. Subconfluent MDA-MB-231 and MCF-7 cells had 2.7- and 2.3-fold higher levels of PPAR mRNA, respectively, compared with that present after 72 h serum deprivation.
between 0.01 and 1 ng total RNA for 18S rRNA (Fig. 2). RNA levels were used that were within these ranges for all subsequent experiments. 3.3. Expression changes in PPARβ in MCF-7, MDA-MB-231 and MCF-10A breast epithelial cell lines The effect of cellular confluence and FBS deprivation on PPAR mRNA levels was examined in MCF-7, MDA-MB-231 and MCF-10A cells using RNA isolated from subconfluent proliferating cells, confluent cells and cells deprived of FBS for 72 h (Fig. 3). Statistical comparisons were made only between cells lines after 72 h FBS deprivation and within cell lines between subconfluent proliferating cells and cells deprived of FBS for 72 h. PPAR mRNA expression was 4.2- and 3.8-fold greater (P < 0.05) in the MDA-MB-231 and MCF-10A cells, respectively, after 72 h FBS deprivation compared with MCF-7 cells (Fig. 3). Within cell
lines, MDA-MB-231 and MCF-7 cells had 2.7- and 2.3-fold greater (P < 0.05) PPAR mRNA expression, respectively, in subconfluent proliferating cells compared with that present after 72 h FBS deprivation (Fig. 3). The non-tumourigenic MCF-10A cell line showed no difference between PPAR levels in subconfluent cells and those deprived of FBS. 3.4. Effect of bezafibrate on PPARβ expression in MCF-7 cells To determine if PPAR mRNA expression could be upregulated by its ligands the lowest PPAR expressing cell line, MCF-7, was chosen for further studies. The addition of the PPAR ligand bezafibrate (50 M) to MCF-7 cells for 24 h resulted in a significant (P < 0.05) increase in PPAR mRNA levels of 1.4-fold compared with cells cultured in the absence of bezafibrate (Fig. 4). Bezafibrate is a clinically used drug used in the treatment of individuals with altered triglyceride levels [29].
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Fig. 4. PPAR mRNA expression in MCF-7 cells in the presence of bezafibrate (50 M). PPAR expression was assessed using real time RT-PCR with reference to 18S rRNA. Results are presented as −CT where CT values were normalised to MCF-7 cells in serum free media in the absence of bezafibrate. Each bar represents mean ± S.E.M. (n = 25–27). Symbol (∗) denotes significant difference. The addition of bezafibrate to MCF-7 cells resulted in a 1.4-fold increase in PPAR mRNA levels.
4. Discussion In this paper, we describe the first study examining PPAR mRNA expression in human breast epithelial cell lines and the effect of proliferation and agonist stimulation on expression patterns. We detected PPAR mRNA in each of the tested human mammary gland epithelial cells providing further evidence of its ubiquitous distribution. This study is the first time PPAR has been identified in human breast epithelial cells and is in agreement with studies describing expression of PPAR in murine mammary gland [21] and other epithelial cells including human tracheobronchial and keratinocyte epithelial cells [22]. The ubiquitous nature of PPAR expression has precluded assumptions to be made about specific physiological roles for this subtype. However, a body of evidence now suggests it has a variety of roles including a function in cell cycle regulation [20], adipocyte differentiation [30,31], colon cancer [17,19] and mouse embryo implantation and decidualisation [11]. Given its wide spread nature PPAR is likely to be pivotal in basic cellular function.
PPAR expression was consistently higher in subconfluent proliferating cells from the tumorigenic MDA-MB-231 and MCF-7 cell lines compared with those that had been deprived of FBS for 72 h, relative to 18S rRNA levels. A similar dynamic expression was not observed for the non-tumorigenic MCF-10A cells, which correlated with the slower growth rate of this cell line. PPAR mRNA expression levels have previously been linked to changes in confluence and differentiation in human foreskin epidermal keratinocytes [22] and these results suggest that PPAR expression may be linked to cell cycle changes in breast epithelial cells. Further evidence for the association between PPAR and cell cycle is highlighted by studies using the PPAR-null mouse where dermal hyperplasia following topical application of 12-O-tetradecanoyl-phorbol-13-acetate results in elevated mRNA expression of cell cycle regulatory genes [20]. MCF-7 cells showed the lowest level of PPAR expression and no correlation was seen between PPAR mRNA levels and tumorigenicity, demonstrating that PPAR mRNA overexpression, as has been reported in primary colon cancers [19], is not an inherent property of tumorigenic human breast epithelial cell
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lines. However, a rigorous study using primary human breast cancer tissue needs to be conducted to ascertain the role of PPAR definitively in this tumour type. Despite expressing a low basal level of PPAR, MCF-7 cells were responsive to the PPAR ligand bezafibrate resulting in a significantly increased level of mRNA. As PPAR mRNA levels are regulated in response to nutritional state in the liver and kidney [7] further studies will be necessary to determine whether PPAR is induced in the mammary gland by fatty acids. This may be important given the promotional effect of dietary fatty acids on mammary gland tumorigenesis [32]. A similar effect was observed in cultured rat pancreatic islets where PPAR␣ mRNA levels were significantly increased in response to the PPAR␣ activator Wy-14,643 [33]. A PPAR response element has recently been described [19] and it appears that the liver fatty-acid-binding protein while regulated by PPAR␣ in the liver is regulated by PPAR in the small intestine [34] despite the presence of PPAR␣ in these cells [35]. Human breast epithelial cells also express PPAR␥ [36–38] and with overlapping substrate specificity the overall response of the cells to PPAR endogenous activators, such as fatty acids, is likely to be complex. In summary, PPAR is expressed in human breast epithelial cell lines. Dynamic changes in mRNA level were seen with culturing in the tumorigenic cell lines. We also demonstrated that PPAR expression is modulated in the presence of its ligand. Since PPAR is postulated to play a role in lipid homeostasis [39] it is likely that it has a functional significance in the mammary gland. Certainly the overexpression of PPAR seen in colon cancer [19] does not appear to be a general feature of human mammary gland tumorigenesis. However, our finding of PPAR expression in breast epithelial cell lines and its response to PPAR activating ligands indicates that this subtype may be significant in mammary gland tumorigenesis through its activation by known tumour promoters such as linoleic acid and through increased expression of cell cycle regulatory genes.
Acknowledgements This work was funded by the National Health and Medical Research Committee grant # 102422. We
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wish to thank Greg Monteith for his helpful discussions regarding this manuscript.
References [1] S. Green, W. Wahli, Peroxisome proliferator-activated receptors: finding the orphan a home, Mol. Cell Endocrinol. 100 (1994) 149–153. [2] A unified nomenclature system for the nuclear receptor superfamily, Cell 97 (1999) 161–163. [3] L. Fajas, D. Auboeuf, E. Raspe, K. Schoonjans, A.M. Lefebvre, R. Saladin, J. Najib, M. Laville, J.C. Fruchart, S. Deeb, A. Vidal-Puig, J. Flier, M.R. Briggs, B. Staels, H. Vidal, J. Auwerx, The organisation, promoter analysis, and expression of the human PPARγ gene, J. Biol. Chem. 272 (1997) 18779–18789. [4] L. Fajas, J. Fruchart, J. Auwerx, PPAR␥3 mRNA: a distinct PPAR␥ mRNA subtype transcribed from an independent promoter, FEBS Lett. 438 (1998) 55–60. [5] T.M. Willson, P.J. Brown, D.D. Sternbach, B.R. Henke, The PPARs: from orphan receptors to drug discovery, J. Med. Chem. 43 (2000) 527–550. [6] J. Skogsberg, K. Kannisto, L. Roshani, E. Gagne, A. Hamsten, C. Larsson, E. Ehrenborg, Characterisation of the human peroxisome proliferator activated receptor δ gene and its expression, Int. J. Mol. Med. 6 (2000) 73–81. [7] P. Escher, O. Braissant, S. Basu-Modak, L. Michalik, W. Wahli, B. Desvergne, Rat PPARs: quantitative analysis in adult rat tissues and regulation in fasting and refeeding, Endocrinology 142 (2001) 4195–4202. [8] B.M. Forman, J. Chen, R.M. Evans, Hypolipidemic drugs, polyunsaturated fatty acids, and eicosanoids are ligands for peroxisome proliferator-activated receptors ␣ and ␦, Proc. Natl. Acad. Sci. U.S.A. 94 (1997) 4312–4317. [9] S.A. Kliewer, S.S. Sundseth, S.A. Jones, P.J. Brown, G.B. Wisely, C.S. Koble, P. Devchand, W. Wahli, T.M. Willson, J.M. Lenhard, J.M. Lehmann, Fatty acids and eicosanoids regulate gene expression through direct interactions with peroxisome proliferator-activated receptors ␣ and ␥, Proc. Natl. Acad. Sci. U.S.A. 94 (1997) 4318–4323. [10] G. Krey, O. Braissant, F. L’Horset, E. Kalkhoven, M. Perroud, M.G. Parker, W. Wahli, Fatty acids, eicosanoids, and hypolipidemic agents identified as ligands of peroxisome proliferator-activated receptors by coactivator-dependent receptor ligand assay, Mol. Endocrinol. 11 (1997) 779–791. [11] H. Lim, R.A. Gupta, W. Ma, B.C. Paria, D.E. Moller, J.D. Morrow, R.N. DuBois, J.M. Trzaskos, S.K. Dey, Cyclo-oxygenase-2-derived prostacyclin mediates embryo implantation in the mouse via PPAR␦, Genes Dev. 13 (1999) 1561–1574. [12] H. Mano, C. Kimura, Y. Fujisawa, T. Kameda, M. Watanabe-Mano, H. Kaneko, T. Kaneda, Y. Hakeda, M. Kumegawa, Cloning and function of rabbit peroxisome proliferator-activated receptor ␦/ in mature osteoclasts, J. Biol. Chem. 275 (2000) 8126–8132.
1058
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[13] S. Basu-Modak, O. Braissant, P. Escher, B. Desvergne, P. Honegger, W. Wahli, Peroxisome proliferator-activated receptor  regulates acyl-CoA synthetase 2 in reaggregated rat brain cell cultures, J. Biol. Chem. 274 (1999) 35881–35888. [14] M.D. Leibowitz, C. Fievet, N. Hennuyer, J. Peinado-Onsurbe, H. Duez, J. Bergera, C.A. Cullinan, C.P. Sparrow, J. Baffic, G.D. Berger, C. Santini, R.W. Marquis, R.L. Tolman, R.G. Smith, D.E. Moller, J. Auwerx, Activation of PPAR␦ alters lipid metabolism in db/db mice, FEBS Lett. 473 (2000) 333– 336. [15] W.R. Oliver Jr., J.L. Shenk, M.R. Snaith, C.S. Russell, K.D. Plunket, N.L. Bodkin, M.C. Lewis, D.A. Winegar, M.L. Sznaidman, M.H. Lambert, H.E. Xu, D.D. Sternbach, S.A. Kliewer, B.C. Hansen, T.M. Willson, A selective peroxisome proliferator-activated receptor ␦ agonist promotes reverse cholesterol transport, Proc. Natl. Acad. Sci. U.S.A. 98 (2001) 5306–5311. [16] L. Michalik, B. Desvergne, N.S. Tan, S. Basu-Modak, P. Escher, J. Rieusset, J.M. Peters, G. Kaya, F.J. Gonzalez, J. Zakany, D. Metzger, P. Chambon, D. Duboule, W. Wahli, Impaired skin wound healing in peroxisome proliferator-activated receptor PPAR␣ and PPAR mutant mice, J. Cell Biol. 154 (2001) 799–814. [17] R.A. Gupta, J. Tan, W.F. Krause, M.W. Geraci, T.M. Willson, S.K. Dey, R.N. DuBois, Prostacyclin-mediated activation of peroxisome proliferator-activated receptor ␦ in colorectal cancer, Proc. Natl. Acad. Sci. U.S.A. 97 (2000) 13275–13280. [18] B.H. Park, B. Vogelstein, K.W. Kinzler, Genetic disruption of PPAR␦ decreases the tumorigenicity of human colon cancer cells, Proc. Natl. Acad. Sci. U.S.A. 98 (2001) 2598–2603. [19] T.C. He, T.A. Chan, B. Vogelstein, K.W. Kinzler, PPAR␦ is an APC-regulated target of nonsteroidal anti-inflammatory drugs, Cell 99 (1999) 335–345. [20] J.M. Peters, S.S.T. Lee, W. Li, J.M. Ward, O. Gavrilova, C. Everett, M.L. Reitman, L.D. Hudson, F.J. Gonzalez, Growth, adipose, brain, and skin alterations resulting from targeted disruption of the mouse peroxisome proliferator-activated receptor  (␦), Mol. Cell Biol. 20 (2000) 5119–5128. [21] J.M. Gimble, G.M. Pighetti, M.R. Lerner, X. Wu, S.A. Lightfoot, D.J. Brackett, K. Darcy, A.B. Hollingsworth, Expression of peroxisome proliferator activated receptor mRNA in normal and tumorigenic rodent mammary glands, Biochem. Biophys. Res. Commun. 253 (1998) 813–817. [22] H. Matsuura, H. Adachi, R.C. Smart, X. Xu, J. Arata, A.M. Jetten, Correlation between expression of peroxisome proliferator-activated receptor  and squamous differentiation in epidermal and tracheobronchial epithelial cells, Mol. Cell Endocrinol. 147 (1999) 85–92. [23] K.H. Hecker, K.H. Roux, High and low annealing temperatures increase both specificity and yield in touchdown and stepdown PCR, Bio. Tech. 20 (1996) 478–485. [24] D.A. Favy, S. Lafarge, P. Rio, C. Vissac, Y.J. Bignon, D. Bernard-Gallon, Real-time PCR quantification of full-length and exon 11 spliced BRCA1 transcripts in human breast cancer cell lines, Biochem. Biophys. Res. Commun. 274 (2000) 73–78.
[25] A.C. White, J.A. Levy, C.M. McGrath, Site-selective growth of a hormone-responsive human breast carcinoma in athymic mice, Cancer Res. 42 (1982) 906–912. [26] J.E. Price, A. Polyzos, R.D. Zhang, L.M. Daniels, Tumorigenicity and metastasis of human breast carcinoma cell lines in nude mice, Cancer Res. 50 (1990) 717–721. [27] W.M. Freeman, S.J. Walker, K.E. Vrana, Quantitative RT-PCR: pitfalls and potential, Bio. Tech. 26 (1999) 112–125. [28] L. Overbergh, D. Valckx, M. Waer, C. Mathieu, Quantification of murine cytokine mRNAs using real time quantitative reverse transcriptase-PCR, Cytokine 11 (1999) 305–312. [29] G.F. Watts, Treating patients with low high-density lipoprotein cholesterol: choices, issues and opportunities, Curr. Control. Trials. Cardiovasc. Med. 2 (2001) 118–122. [30] C. Bastie, S. Luquet, D. Holst, C. Jehl-Pietri, P.A. Grimaldi, Alterations of peroxisome proliferator-activated receptor ␦ activity affect fatty acid-controlled adipose differentiation, J. Biol. Chem. 275 (2000) 38768–38773. [31] C. Bastie, D. Holst, D. Gaillard, C. Jehl-Pietri, P.A. Grimaldi, Expression of peroxisome proliferator-activated receptor PPAR␦ promotes induction of PPAR␥ and adipocyte differentiation in 3T3C2 fibroblasts, J. Biol. Chem. 274 (1999) 21920–21925. [32] J.H. Weisburger, Dietary fat and risk of chronic disease: mechanistic insights from experimental studies, J. Am. Diet. Assoc. 97 (1997) S16–23. [33] H. Yoshikawa, Y. Tajiri, Y. Sako, T. Hashimoto, F. Umeda, H. Nawata, Effects of bezafibrate on  cell function of rat pancreatic islets, Eur. J. Pharmacol. 426 (2001) 201–206. [34] H. Poirier, I. Niot, M.C. Monnot, O. Braissant, C. Meunier-Durmort, P. Costet, T. Pineau, W. Wahli, T.M. Willson, P. Besnard, Differential involvement of peroxisomeproliferator-activated receptors ␣ and ␦ in fibrate and fatty-acid-mediated inductions of the gene encoding liver fatty-acid-binding protein in the liver and the small intestine, Biochem. J. 355 (2001) 481–488. [35] O. Braissant, F. Foufelle, C. Scotto, M. Daucu, W. Wahli, Differential expression of peroxisome proliferator-activated receptors (PPARs): tissue distribution of PPAR␣, -, and -␥ in the adult rat, Endocrinology 137 (1996) 354–366. [36] E. Mueller, P. Sarraf, P. Tontonoz, R. Evans, K. Martin, M. Zhang, C. Fletcher, S. Singer, B. Spiegelman, Terminal differentiation of human breast cancer through PPAR␥, Mol. Cell 1 (1998) 465–470. [37] M.W. Kilgore, P.L. Tate, S. Rai, E. Sengoku, T.M. Price, MCF-7 and T47D human breast cancer cells contain a functional peroxisomal response, Mol. Cell Endocrinol. 129 (1997) 229–235. [38] E. Elstner, C. Muller, K. Koshizuka, E.A. Williamson, D. Park, H. Asou, P. Shintaku, J.W. Said, D. Heber, H.P. Koeffler, Ligands for peroxisome proliferator-activated receptor ␥ and retinoic acid receptor inhibit growth and induce apoptosis of human breast cancer cells in vitro and in BNX mice, Proc. Natl. Acad. Sci. U.S.A. 95 (1998) 8806–8811. [39] J.P.V. Heuvel, J.M. Peters, Biology of the PPAR family of receptors, Comm. Toxicol. 7 (2001) 233–258.
Peroxisome proliferator-activated receptor  expression in human breast epithelial cell lines of tumorigenic and non-tumorigenic origin Kate M. Suchanek, Fiona J. May, Won Jae Lee, Nicola A. Holman, Sarah J. Roberts-Thomson∗ School of Pharmacy, The University of Queensland, St. Lucia, QLD 4072, Australia Received 23 November 2001; received in revised form 6 February 2002; accepted 7 February 2002
Abstract Peroxisome proliferator-activated receptor  (PPAR) is a member of the nuclear hormone receptor superfamily and is a ligand activated transcription factor, although the precise genes that it regulates and its physiological and pathophysiological role remain unclear. In view of the association of PPAR with colon cancer and increased mRNA levels of PPAR in colon tumours we sought in this study to examine the expression of PPAR in human breast epithelial cells of tumorigenic (MCF-7 and MDA-MB-231) and non-tumorigenic origin (MCF-10A). Using quantitative RT-PCR we measured PPAR mRNA levels in MCF-7, MDA-MB-231 and MCF-10A cells at various stages in culture. After serum-deprivation, MDA-MB-231 and MCF-10A cells had a 4.2- and 3.8-fold statistically greater expression of PPAR compared with MCF-7 cells. The tumorigenic cell lines also exhibited a significantly greater level of PPAR mRNA after serum deprivation compared with subconfluence whereas such an effect was not observed in non-tumorigenic MCF-10A cells. The expression of PPAR was inducible upon exposure to the PPAR ligand bezafibrate. Our results suggest that unlike colon cancer, PPAR overexpression is not an inherent property of breast cancer cell lines. However, the dynamic changes in PPAR mRNA expression and the ability of PPAR in the MCF-7 cells to respond to ligand indicates that PPAR may play a role in mammary gland carcinogenesis through activation of downstream genes via endogenous fatty acid ligands or exogenous agonists. © 2002 Elsevier Science Ltd. All rights reserved. Keywords: PPAR; Breast; Human; Cell lines
1. Introduction Abbreviations: ATCC, American type culture collection; CT , threshold cycle; DMEM, Dulbecco’s modified eagle medium; FBS, fetal bovine serum; Real time RT-PCR, real time reverse transcriptase-PCR; PPAR, peroxisome proliferator-activated receptor; RTPCR, reverse transcriptase-PCR; S.E.M., standard error of the mean ∗ Corresponding author. Tel.: +61-7-3365-3193; fax: +61-7-3365-1688. E-mail address: [email protected] (S.J. Roberts-Thomson).
Peroxisome proliferator-activated receptor  (PPAR; NR1C2) is a member of the nuclear hormone receptor superfamily that includes the thyroid hormone, Vitamin D and retinoic acid receptors [1]. Members of the PPAR family also include PPAR␣, PPAR␥1, PPAR␥2 and PPAR␥3 [2–4]. While PPAR␣ and PPAR␥ have clearly defined roles
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in controlling lipid and glucose homeostasis (reviewed in [5]) the physiological role of PPAR remains less defined with elucidation of its function hindered by a lack of a widely available specific agonist. PPAR is expressed in humans at similar levels in all tissues so far examined [6]. Its expression in rodent tissues is also ubiquitous although its expression level is often greater than that of PPAR␣ or ␥ [7]. Endogenous ligands of PPAR include the polyunsaturated fatty acids arachidonic acid and linoleic acid [8], which also act as agonists for PPAR␣ and ␥ [8–10]. PPAR has been associated with a number of biological functions including embryo implantation [11], osteoclastic bone resorption [12], lipid homeostasis [13–15], skin wound healing [16] and colon cancer [17–19]. Using PPAR-null mice in vivo evidence has been presented linking PPAR with roles in development, myelination of the corpus callosum, lipid metabolism and epidermal cell proliferation [20]. Significantly, studies using the PPAR-null mice [20] highlight a role for PPAR in cell cycle control, a finding that supports an earlier study suggesting a link between PPAR and human colon cancer [19]. In four individual patients with colorectal cancer, increased PPAR mRNA expression was seen in cancer tissue compared with normal colon epithelium, suggesting that the level of PPAR expression may be a factor influencing colon carcinogenesis [19]. Indeed, a total absence of PPAR in a human colorectal cancer cell line reduced the ability of the cells to form tumours in nude mice [18]. PPAR is expressed in the mammary gland of the mouse [21] as well as cells of epithelial origin [22]. Despite the ubiquitous nature of PPAR expression and its association with tumorigenesis, there are no studies examining the expression of PPAR in human breast epithelial cells. We have sought in this study to examine the expression of PPAR in two human breast cancer epithelial cell lines and a non-tumorigenic breast epithelial cell line and assess changes in PPAR expression levels during proliferation and in response to agonist stimulation as a first step in characterising any potential role for this receptor in mammary gland tumorigenesis.
2. Materials and methods 2.1. Materials Cell culture media, antibiotics and fetal bovine serum (FBS) were obtained from Life Technologies (Mulgrave, Australia). MCF-7, MDA-MB-231 and MCF-10A cell lines were purchased from the American type culture collection (ATCC). Six well plates were purchased from Nalge Nunc International (IL, USA) and bezafibrate was purchased from Sigma (Castle Hill, Australia). The human brain cDNA library was the kind gift of Professor M.E. McManus (The University of Queensland, Australia). All other reagents and materials were purchased from Sigma. 2.2. Cell culture Cells were cultured in Dulbecco’s modified eagle medium (DMEM) supplemented with 10% FBS, glutamine (2 mM), penicillin G (100 units/ml) and streptomycin sulphate (100 mg/ml) in 6 well plates and maintained at 37 ◦ C and 5% CO2 . Prior to confluence, cells were trypsinised with a 0.05% trypsin/0.53 mM EDTA solution and resuspended in fresh growth media before plating, at a ratio of 1:6, onto a new growth surface. RNA was isolated from subconfluent proliferating cells and confluent cells (>95% confluence). For studies involving FBS deprivation, confluent cells were cultured for 72 h in FBS-free media prior to RNA isolation. For studies involving the addition of bezafibrate (50 M) MCF-7 cells were allowed to reach confluence in normal growth media then cultured in the absence of FBS for 48 h prior to the addition of bezafibrate (in DMSO) in fresh FBS-free media for 24 h. Control cells were incubated in the presence of DMSO 0.12%. All experiments were performed in triplicate. 2.3. RNA isolation The QIAGEN RNeasyTM mini kit (QIAGEN, Clifton Hill, Vic., Australia) was used to isolate total RNA and included an on-column DNase treatment (QIAGEN). RNA was resuspended in RNase free water and stored at −80 ◦ C until required. RNA was quantitated by absorbance at 260 nm while purity was determined by the ratio of absorbance at 260 and 280 nm.
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2.4. Reverse transcriptase-PCR The access RT-PCR system (Promega, Sydney, Australia) was used for reverse transcriptase-PCR (RT-PCR). Following reverse transcription at 48 ◦ C for 45 min using gene-specific primers a Stepdown PCR protocol was followed [23]. The annealing temperature ranged from 65 to 49 ◦ C for 30 s with extension at 68 ◦ C for 30 s. Each reaction contained 0.5 g RNA, Mg2+ (1.5 mM for MDA-MB-231 and brain cDNA library; 2 mM for MCF-7 and -10A) primers 0.5 M and dNTPs 0.2 mM. Primers for human PPAR were as follows; forward: 5 -AAAGAAGGCCCGCAGCAT3 , reverse: 5-CCTTCTCTGCCTGCCACAAT-3 and were designed to amplify a 92 bp product. For all reactions a human brain cDNA library was used as a positive control and reactions run in the absence of the reverse transcriptase enzyme were used as negative controls. RT-PCR reactions were analysed by agarose gel electrophoresis and bands were excised from the gel and sequenced to confirm identity. 2.5. Real time RT-PCR Primers and probes for real time RT-PCR were designed using the software Primer Express Version 1.0 (Applied Biosystems, Foster City, USA). Primers for PPAR were as described above. The probe (5 -TGCTGCGCTGCACCCGCT-3 ) had the fluorescent dye FAM (6-carboxy-fluorescin) attached to the 5 end and the quencher dye TAMRA (6-carboxy-tetramethylrhodamine) at the 3 end. Probe and primers for the standard 18S rRNA were purchased from Applied Biosystems. RNA isolated from MCF-7 cells cultured in normal growth media was used to obtain the standard curve using 0.1–100 ng total RNA for PPAR amplification and 0.001–1 ng total RNA for 18S rRNA amplification. Equal amplification efficiency of the target and standard amplicons was determined by linear regression analysis and examination of R2 using Microsoft® Excel (Microsoft Corporation, Washington, USA). Using the Taqman® Gold RT-PCR kit (Applied Biosystems), each real time RT-PCR reaction (25 ml) contained 1 × Taqman A buffer, 5.5 mM magnesium chloride, 300 mM dATP, dCTP and dGTP, 600 mM dUTP, AmpliTaq Gold DNA polymerase 0.625 units, MultiScribe reverse transcriptase 0.625 units, forward
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and reverse primers (PPAR 900 nM, 18S rRNA 50 nM), RNA 25 ng (PPAR) or 0.25 ng (18S rRNA), probe (PPAR 200 nM, 18S rRNA 50 nM) and RNase inhibitor 10 units. RNA was replaced with RNase free water in no template controls. Reactions were cycled in an ABI PRISM 7700 Sequence Detector (Applied Biosystems) using the following conditions; 48 ◦ C for 30 min, 95 ◦ C for 10 min then 45 cycles of 95 ◦ C for 15 s and 60 ◦ C for 1 min, followed by 25 ◦ C for 10 min. For real time RT-PCR CT values (cycle number at which amplicon product is first detected above baseline) were used to calculate CT for each sample (CT PPAR␣ − C T 18S rRNA). To allow comparison between cell lines, a CT value was calculated by normalising values to MDA-MB-231, 72 h FBS deprivation. The MDA-MB-231 cell line was chosen for normalisation due to its greater levels of PPAR mRNA compared with MCF-10A and -7 cells cultured under the same conditions. The 72 h time point was chosen, as this time point was used to compare cell lines. Fold change is calculated from CT using the formula 2−CT [24]. Statistical analysis of samples was performed using SigmaStat® (SPSS Science, Chicago, IL, USA). Results were analysed using oneway ANOVA with the Tukey Test for all pairwise multiple comparison for normally distributed data of equal variance, or where data were not normally distributed the Kruskal–Wallis ANOVA on Ranks with Dunn’s Method of Multiple Comparison was used. Significant (P < 0.05) values are indicated in the result section.
3. Results 3.1. Expression of PPARβ in breast epithelial cell lines Using RT-PCR we showed that PPAR is expressed in human breast epithelial cell lines (Fig. 1): tumorigenic MCF-7 and MDA-MB-231 [25] and non-tumorigenic MCF-10A [26]. 3.2. Development of a quantitative assay for PPARβ mRNA in breast epithelial cell lines A real time RT-PCR assay was developed to quantify the mRNA level of PPAR in relation to 18S rRNA
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Fig. 1. RT-PCR amplification of a 92 bp fragment of PPAR in breast epithelial cell lines. The PPAR fragment was amplified as outlined in Section 2: lane 1, marker; lane 2, human brain cDNA control; lane 3, MDA-MB-231; lane 4, MDA-MB-231, -reverse transcriptase (RT); lane 5, MCF-7; lane 6, MCF-7, -RT; lane 7, MCF-10A; lane 8, MCF-10A, -RT.
levels. We initially validated the real time RT-PCR assay to ensure that our target and internal standard amplicons were amplified with the same efficiency. Relative comparisons can be made with regard to amplicon levels when efficiencies are equal [27]. To determine this we examined the relationship between log (total RNA) and CT values (CT values are defined as the fraction cycle number at which fluorescence passes
a predetermined threshold (set at 10 times the standard deviation of the baseline [28]) for the amplification of PPAR and 18S rRNA (Fig. 2). If the slope of a plot of log (input amount) versus CT (CT PPAR − C T 18S rRNA) has an absolute value of <0.1 this suggests an approximate equal amplification efficiency. We determined that equal amplification efficiency lies between 1 and 100 ng total RNA for PPAR and
Fig. 2. Standard curve for the real time RT-PCR amplification of PPAR and 18S rRNA in MCF-7 human breast cancer epithelial cells. Each data point represents the mean ± S.D. (n = 3). The x-axis shows the concentration of total RNA on a logarithmic scale. The RNA was diluted 100-fold for the measurement of 18S rRNA.
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Fig. 3. PPAR mRNA expression in MDA-MB-231, MCF-7 and -10A human breast epithelial cell lines by real time RT-PCR with reference to 18S rRNA. Expression levels were quantitated in subconfluent proliferating cells and at >95% confluence (confluent cells) and after 72 h in FBS deprivation as described in Section 2. CT have been normalised to MDA-MB-231, 72 h FBS deprivation to give a CT value for each group. The MDA-MB-231 cell line was chosen for normalisation due to its greater levels of PPAR mRNA compared with MCF-10A and -7 cells cultured under the same conditions. Each bar represents mean ± S.E.M. (n: subconfluence, confluence, 72 h FBS deprivation; MDA-MB-231 and MCF-10A; 12, 21, 33; MCF-7, 12, 11, 22, respectively). Symbol (∗) denotes significant difference. After 72 h serum deprivation, MDA-MB-231 and MCF-10A cells had 4.2- and 3.8-fold higher levels of PPAR mRNA, respectively, than MCF-7 cells. Subconfluent MDA-MB-231 and MCF-7 cells had 2.7- and 2.3-fold higher levels of PPAR mRNA, respectively, compared with that present after 72 h serum deprivation.
between 0.01 and 1 ng total RNA for 18S rRNA (Fig. 2). RNA levels were used that were within these ranges for all subsequent experiments. 3.3. Expression changes in PPARβ in MCF-7, MDA-MB-231 and MCF-10A breast epithelial cell lines The effect of cellular confluence and FBS deprivation on PPAR mRNA levels was examined in MCF-7, MDA-MB-231 and MCF-10A cells using RNA isolated from subconfluent proliferating cells, confluent cells and cells deprived of FBS for 72 h (Fig. 3). Statistical comparisons were made only between cells lines after 72 h FBS deprivation and within cell lines between subconfluent proliferating cells and cells deprived of FBS for 72 h. PPAR mRNA expression was 4.2- and 3.8-fold greater (P < 0.05) in the MDA-MB-231 and MCF-10A cells, respectively, after 72 h FBS deprivation compared with MCF-7 cells (Fig. 3). Within cell
lines, MDA-MB-231 and MCF-7 cells had 2.7- and 2.3-fold greater (P < 0.05) PPAR mRNA expression, respectively, in subconfluent proliferating cells compared with that present after 72 h FBS deprivation (Fig. 3). The non-tumourigenic MCF-10A cell line showed no difference between PPAR levels in subconfluent cells and those deprived of FBS. 3.4. Effect of bezafibrate on PPARβ expression in MCF-7 cells To determine if PPAR mRNA expression could be upregulated by its ligands the lowest PPAR expressing cell line, MCF-7, was chosen for further studies. The addition of the PPAR ligand bezafibrate (50 M) to MCF-7 cells for 24 h resulted in a significant (P < 0.05) increase in PPAR mRNA levels of 1.4-fold compared with cells cultured in the absence of bezafibrate (Fig. 4). Bezafibrate is a clinically used drug used in the treatment of individuals with altered triglyceride levels [29].
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Fig. 4. PPAR mRNA expression in MCF-7 cells in the presence of bezafibrate (50 M). PPAR expression was assessed using real time RT-PCR with reference to 18S rRNA. Results are presented as −CT where CT values were normalised to MCF-7 cells in serum free media in the absence of bezafibrate. Each bar represents mean ± S.E.M. (n = 25–27). Symbol (∗) denotes significant difference. The addition of bezafibrate to MCF-7 cells resulted in a 1.4-fold increase in PPAR mRNA levels.
4. Discussion In this paper, we describe the first study examining PPAR mRNA expression in human breast epithelial cell lines and the effect of proliferation and agonist stimulation on expression patterns. We detected PPAR mRNA in each of the tested human mammary gland epithelial cells providing further evidence of its ubiquitous distribution. This study is the first time PPAR has been identified in human breast epithelial cells and is in agreement with studies describing expression of PPAR in murine mammary gland [21] and other epithelial cells including human tracheobronchial and keratinocyte epithelial cells [22]. The ubiquitous nature of PPAR expression has precluded assumptions to be made about specific physiological roles for this subtype. However, a body of evidence now suggests it has a variety of roles including a function in cell cycle regulation [20], adipocyte differentiation [30,31], colon cancer [17,19] and mouse embryo implantation and decidualisation [11]. Given its wide spread nature PPAR is likely to be pivotal in basic cellular function.
PPAR expression was consistently higher in subconfluent proliferating cells from the tumorigenic MDA-MB-231 and MCF-7 cell lines compared with those that had been deprived of FBS for 72 h, relative to 18S rRNA levels. A similar dynamic expression was not observed for the non-tumorigenic MCF-10A cells, which correlated with the slower growth rate of this cell line. PPAR mRNA expression levels have previously been linked to changes in confluence and differentiation in human foreskin epidermal keratinocytes [22] and these results suggest that PPAR expression may be linked to cell cycle changes in breast epithelial cells. Further evidence for the association between PPAR and cell cycle is highlighted by studies using the PPAR-null mouse where dermal hyperplasia following topical application of 12-O-tetradecanoyl-phorbol-13-acetate results in elevated mRNA expression of cell cycle regulatory genes [20]. MCF-7 cells showed the lowest level of PPAR expression and no correlation was seen between PPAR mRNA levels and tumorigenicity, demonstrating that PPAR mRNA overexpression, as has been reported in primary colon cancers [19], is not an inherent property of tumorigenic human breast epithelial cell
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lines. However, a rigorous study using primary human breast cancer tissue needs to be conducted to ascertain the role of PPAR definitively in this tumour type. Despite expressing a low basal level of PPAR, MCF-7 cells were responsive to the PPAR ligand bezafibrate resulting in a significantly increased level of mRNA. As PPAR mRNA levels are regulated in response to nutritional state in the liver and kidney [7] further studies will be necessary to determine whether PPAR is induced in the mammary gland by fatty acids. This may be important given the promotional effect of dietary fatty acids on mammary gland tumorigenesis [32]. A similar effect was observed in cultured rat pancreatic islets where PPAR␣ mRNA levels were significantly increased in response to the PPAR␣ activator Wy-14,643 [33]. A PPAR response element has recently been described [19] and it appears that the liver fatty-acid-binding protein while regulated by PPAR␣ in the liver is regulated by PPAR in the small intestine [34] despite the presence of PPAR␣ in these cells [35]. Human breast epithelial cells also express PPAR␥ [36–38] and with overlapping substrate specificity the overall response of the cells to PPAR endogenous activators, such as fatty acids, is likely to be complex. In summary, PPAR is expressed in human breast epithelial cell lines. Dynamic changes in mRNA level were seen with culturing in the tumorigenic cell lines. We also demonstrated that PPAR expression is modulated in the presence of its ligand. Since PPAR is postulated to play a role in lipid homeostasis [39] it is likely that it has a functional significance in the mammary gland. Certainly the overexpression of PPAR seen in colon cancer [19] does not appear to be a general feature of human mammary gland tumorigenesis. However, our finding of PPAR expression in breast epithelial cell lines and its response to PPAR activating ligands indicates that this subtype may be significant in mammary gland tumorigenesis through its activation by known tumour promoters such as linoleic acid and through increased expression of cell cycle regulatory genes.
Acknowledgements This work was funded by the National Health and Medical Research Committee grant # 102422. We
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wish to thank Greg Monteith for his helpful discussions regarding this manuscript.
References [1] S. Green, W. Wahli, Peroxisome proliferator-activated receptors: finding the orphan a home, Mol. Cell Endocrinol. 100 (1994) 149–153. [2] A unified nomenclature system for the nuclear receptor superfamily, Cell 97 (1999) 161–163. [3] L. Fajas, D. Auboeuf, E. Raspe, K. Schoonjans, A.M. Lefebvre, R. Saladin, J. Najib, M. Laville, J.C. Fruchart, S. Deeb, A. Vidal-Puig, J. Flier, M.R. Briggs, B. Staels, H. Vidal, J. Auwerx, The organisation, promoter analysis, and expression of the human PPARγ gene, J. Biol. Chem. 272 (1997) 18779–18789. [4] L. Fajas, J. Fruchart, J. Auwerx, PPAR␥3 mRNA: a distinct PPAR␥ mRNA subtype transcribed from an independent promoter, FEBS Lett. 438 (1998) 55–60. [5] T.M. Willson, P.J. Brown, D.D. Sternbach, B.R. Henke, The PPARs: from orphan receptors to drug discovery, J. Med. Chem. 43 (2000) 527–550. [6] J. Skogsberg, K. Kannisto, L. Roshani, E. Gagne, A. Hamsten, C. Larsson, E. Ehrenborg, Characterisation of the human peroxisome proliferator activated receptor δ gene and its expression, Int. J. Mol. Med. 6 (2000) 73–81. [7] P. Escher, O. Braissant, S. Basu-Modak, L. Michalik, W. Wahli, B. Desvergne, Rat PPARs: quantitative analysis in adult rat tissues and regulation in fasting and refeeding, Endocrinology 142 (2001) 4195–4202. [8] B.M. Forman, J. Chen, R.M. Evans, Hypolipidemic drugs, polyunsaturated fatty acids, and eicosanoids are ligands for peroxisome proliferator-activated receptors ␣ and ␦, Proc. Natl. Acad. Sci. U.S.A. 94 (1997) 4312–4317. [9] S.A. Kliewer, S.S. Sundseth, S.A. Jones, P.J. Brown, G.B. Wisely, C.S. Koble, P. Devchand, W. Wahli, T.M. Willson, J.M. Lenhard, J.M. Lehmann, Fatty acids and eicosanoids regulate gene expression through direct interactions with peroxisome proliferator-activated receptors ␣ and ␥, Proc. Natl. Acad. Sci. U.S.A. 94 (1997) 4318–4323. [10] G. Krey, O. Braissant, F. L’Horset, E. Kalkhoven, M. Perroud, M.G. Parker, W. Wahli, Fatty acids, eicosanoids, and hypolipidemic agents identified as ligands of peroxisome proliferator-activated receptors by coactivator-dependent receptor ligand assay, Mol. Endocrinol. 11 (1997) 779–791. [11] H. Lim, R.A. Gupta, W. Ma, B.C. Paria, D.E. Moller, J.D. Morrow, R.N. DuBois, J.M. Trzaskos, S.K. Dey, Cyclo-oxygenase-2-derived prostacyclin mediates embryo implantation in the mouse via PPAR␦, Genes Dev. 13 (1999) 1561–1574. [12] H. Mano, C. Kimura, Y. Fujisawa, T. Kameda, M. Watanabe-Mano, H. Kaneko, T. Kaneda, Y. Hakeda, M. Kumegawa, Cloning and function of rabbit peroxisome proliferator-activated receptor ␦/ in mature osteoclasts, J. Biol. Chem. 275 (2000) 8126–8132.
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[13] S. Basu-Modak, O. Braissant, P. Escher, B. Desvergne, P. Honegger, W. Wahli, Peroxisome proliferator-activated receptor  regulates acyl-CoA synthetase 2 in reaggregated rat brain cell cultures, J. Biol. Chem. 274 (1999) 35881–35888. [14] M.D. Leibowitz, C. Fievet, N. Hennuyer, J. Peinado-Onsurbe, H. Duez, J. Bergera, C.A. Cullinan, C.P. Sparrow, J. Baffic, G.D. Berger, C. Santini, R.W. Marquis, R.L. Tolman, R.G. Smith, D.E. Moller, J. Auwerx, Activation of PPAR␦ alters lipid metabolism in db/db mice, FEBS Lett. 473 (2000) 333– 336. [15] W.R. Oliver Jr., J.L. Shenk, M.R. Snaith, C.S. Russell, K.D. Plunket, N.L. Bodkin, M.C. Lewis, D.A. Winegar, M.L. Sznaidman, M.H. Lambert, H.E. Xu, D.D. Sternbach, S.A. Kliewer, B.C. Hansen, T.M. Willson, A selective peroxisome proliferator-activated receptor ␦ agonist promotes reverse cholesterol transport, Proc. Natl. Acad. Sci. U.S.A. 98 (2001) 5306–5311. [16] L. Michalik, B. Desvergne, N.S. Tan, S. Basu-Modak, P. Escher, J. Rieusset, J.M. Peters, G. Kaya, F.J. Gonzalez, J. Zakany, D. Metzger, P. Chambon, D. Duboule, W. Wahli, Impaired skin wound healing in peroxisome proliferator-activated receptor PPAR␣ and PPAR mutant mice, J. Cell Biol. 154 (2001) 799–814. [17] R.A. Gupta, J. Tan, W.F. Krause, M.W. Geraci, T.M. Willson, S.K. Dey, R.N. DuBois, Prostacyclin-mediated activation of peroxisome proliferator-activated receptor ␦ in colorectal cancer, Proc. Natl. Acad. Sci. U.S.A. 97 (2000) 13275–13280. [18] B.H. Park, B. Vogelstein, K.W. Kinzler, Genetic disruption of PPAR␦ decreases the tumorigenicity of human colon cancer cells, Proc. Natl. Acad. Sci. U.S.A. 98 (2001) 2598–2603. [19] T.C. He, T.A. Chan, B. Vogelstein, K.W. Kinzler, PPAR␦ is an APC-regulated target of nonsteroidal anti-inflammatory drugs, Cell 99 (1999) 335–345. [20] J.M. Peters, S.S.T. Lee, W. Li, J.M. Ward, O. Gavrilova, C. Everett, M.L. Reitman, L.D. Hudson, F.J. Gonzalez, Growth, adipose, brain, and skin alterations resulting from targeted disruption of the mouse peroxisome proliferator-activated receptor  (␦), Mol. Cell Biol. 20 (2000) 5119–5128. [21] J.M. Gimble, G.M. Pighetti, M.R. Lerner, X. Wu, S.A. Lightfoot, D.J. Brackett, K. Darcy, A.B. Hollingsworth, Expression of peroxisome proliferator activated receptor mRNA in normal and tumorigenic rodent mammary glands, Biochem. Biophys. Res. Commun. 253 (1998) 813–817. [22] H. Matsuura, H. Adachi, R.C. Smart, X. Xu, J. Arata, A.M. Jetten, Correlation between expression of peroxisome proliferator-activated receptor  and squamous differentiation in epidermal and tracheobronchial epithelial cells, Mol. Cell Endocrinol. 147 (1999) 85–92. [23] K.H. Hecker, K.H. Roux, High and low annealing temperatures increase both specificity and yield in touchdown and stepdown PCR, Bio. Tech. 20 (1996) 478–485. [24] D.A. Favy, S. Lafarge, P. Rio, C. Vissac, Y.J. Bignon, D. Bernard-Gallon, Real-time PCR quantification of full-length and exon 11 spliced BRCA1 transcripts in human breast cancer cell lines, Biochem. Biophys. Res. Commun. 274 (2000) 73–78.
[25] A.C. White, J.A. Levy, C.M. McGrath, Site-selective growth of a hormone-responsive human breast carcinoma in athymic mice, Cancer Res. 42 (1982) 906–912. [26] J.E. Price, A. Polyzos, R.D. Zhang, L.M. Daniels, Tumorigenicity and metastasis of human breast carcinoma cell lines in nude mice, Cancer Res. 50 (1990) 717–721. [27] W.M. Freeman, S.J. Walker, K.E. Vrana, Quantitative RT-PCR: pitfalls and potential, Bio. Tech. 26 (1999) 112–125. [28] L. Overbergh, D. Valckx, M. Waer, C. Mathieu, Quantification of murine cytokine mRNAs using real time quantitative reverse transcriptase-PCR, Cytokine 11 (1999) 305–312. [29] G.F. Watts, Treating patients with low high-density lipoprotein cholesterol: choices, issues and opportunities, Curr. Control. Trials. Cardiovasc. Med. 2 (2001) 118–122. [30] C. Bastie, S. Luquet, D. Holst, C. Jehl-Pietri, P.A. Grimaldi, Alterations of peroxisome proliferator-activated receptor ␦ activity affect fatty acid-controlled adipose differentiation, J. Biol. Chem. 275 (2000) 38768–38773. [31] C. Bastie, D. Holst, D. Gaillard, C. Jehl-Pietri, P.A. Grimaldi, Expression of peroxisome proliferator-activated receptor PPAR␦ promotes induction of PPAR␥ and adipocyte differentiation in 3T3C2 fibroblasts, J. Biol. Chem. 274 (1999) 21920–21925. [32] J.H. Weisburger, Dietary fat and risk of chronic disease: mechanistic insights from experimental studies, J. Am. Diet. Assoc. 97 (1997) S16–23. [33] H. Yoshikawa, Y. Tajiri, Y. Sako, T. Hashimoto, F. Umeda, H. Nawata, Effects of bezafibrate on  cell function of rat pancreatic islets, Eur. J. Pharmacol. 426 (2001) 201–206. [34] H. Poirier, I. Niot, M.C. Monnot, O. Braissant, C. Meunier-Durmort, P. Costet, T. Pineau, W. Wahli, T.M. Willson, P. Besnard, Differential involvement of peroxisomeproliferator-activated receptors ␣ and ␦ in fibrate and fatty-acid-mediated inductions of the gene encoding liver fatty-acid-binding protein in the liver and the small intestine, Biochem. J. 355 (2001) 481–488. [35] O. Braissant, F. Foufelle, C. Scotto, M. Daucu, W. Wahli, Differential expression of peroxisome proliferator-activated receptors (PPARs): tissue distribution of PPAR␣, -, and -␥ in the adult rat, Endocrinology 137 (1996) 354–366. [36] E. Mueller, P. Sarraf, P. Tontonoz, R. Evans, K. Martin, M. Zhang, C. Fletcher, S. Singer, B. Spiegelman, Terminal differentiation of human breast cancer through PPAR␥, Mol. Cell 1 (1998) 465–470. [37] M.W. Kilgore, P.L. Tate, S. Rai, E. Sengoku, T.M. Price, MCF-7 and T47D human breast cancer cells contain a functional peroxisomal response, Mol. Cell Endocrinol. 129 (1997) 229–235. [38] E. Elstner, C. Muller, K. Koshizuka, E.A. Williamson, D. Park, H. Asou, P. Shintaku, J.W. Said, D. Heber, H.P. Koeffler, Ligands for peroxisome proliferator-activated receptor ␥ and retinoic acid receptor inhibit growth and induce apoptosis of human breast cancer cells in vitro and in BNX mice, Proc. Natl. Acad. Sci. U.S.A. 95 (1998) 8806–8811. [39] J.P.V. Heuvel, J.M. Peters, Biology of the PPAR family of receptors, Comm. Toxicol. 7 (2001) 233–258.