The Pennsylvania State University. The Graduate School. The Huck Institutes for Life Sciences MODULATION OF SKIN CANCER BY PEROXISOME

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1 The Pennsylvania State University The Graduate School The Huck Institutes for Life Sciences MODULATION OF SKIN CANCER BY PEROXISOME PROLIFERATOR-ACTIVATED RECEPTOR BETA/DELTA A Dissertation in Molecular Medicine by Bokai Zhu 2012 Bokai Zhu Submitted in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy August 2012

2 The dissertation of Bokai Zhu was reviewed and approved* by the following: Jeffrey M. Peters Distinguished Professor of Environmental Toxicology Dissertation Advisor Chair of Committee Adam B. Glick Associate Professor of Veterinary and Biomedical Sciences Director of the Center for Molecular Toxicology Chair of Molecular Medicine program Gary H. Perdew John T. and Paige S. Smith Professor in Agricultural Sciences Yanming Wang Associate Professor of Biochemistry and Molecular Biology Wendy Hanna-Rose Associate Professor of Biochemistry and Molecular Biology *Signatures are on file in the Graduate School. ii

3 ABSTRACT Ligand activation of peroxisome proliferator activated receptor-β/δ (PPARβ/δ) inhibits chemically-induced skin tumorigenesis and Pparβ/δ-null mice exhibit enhanced chemically-induced skin tumorigenesis compared to wild-type mice. Since over 90% of chemically-induced tumors contain Hras mutations, this suggests that PPARβ/δ could inhibit chemically-induced skin tumorigenesis by modulating Hras signaling, which was examined in this study. In addition, ligand activation of PPARβ/δ and inhibition of cyclooxygenase-2 (COX2) activity by nonsteroidal anti-inflammatory drugs (NSAID) can both attenuate skin tumorigenesis. The hypothesis that combining ligand activation of PPARβ/δ with inhibition of COX2 activity will increase the efficacy of chemoprevention of chemically induced skin tumorigenesis over that observed with either approach alone is also examined in this study. Ligand activation of PPARβ/δ caused a negative selection against cells expressing higher levels of HRAS by inducing a mitotic block. Mitosis-related genes that are predominantly regulated by E2F were induced to a higher level in HRAS-expressing Pparβ/δ-null keratinocytes as compared to HRAS-expressing wild-type keratinocytes. Ligand activated PPARβ/δ repressed expression of these genes by direct binding with p130/p107, facilitating nuclear translocation, and increasing promoter recruitment of p130/p107. In addition, co-treatment with a PPARβ/δ ligand and various mitosis inhibitors increases the efficacy of increasing G2/M arrest. Moreover, PPARβ/δ suppresses tumorigenesis by promoting HRAS-induced senescence. PPARβ/δ transcriptionally regulates Rasgrp1 and Ilk iii

4 causing increased p-erk and decreased p-akt that promote HRAS-induced senescence. PPARβ/δ also promotes senescence through attenuation of HRAS-induced ER stress and ER stress-associated UPR by repressing p-akt-mtor signaling. Further, these studies demonstrate a novel positive feedback loop between p-akt, ER stress and UPR. An acute increase of ER stress is sufficient to establish the positive loop, maintaining higher UPR and p-akt activity, collectively causing evasion of senescence and malignant conversion. Moreover, increased PPARβ/δ expression and decreased ER stress correlated with increased senescence in both mouse and human tumors. Ligand activation of PPARβ/δ with GW0742 caused a PPARβ/δ-dependent delay in the onset of tumor formation. Nimesulide also delayed the onset of tumor formation and caused inhibition of tumor multiplicity in wild-type mice but not in Pparβ/δ-null mice. Combining ligand activation of PPARβ/δ with dietary nimesulide resulted in a further decrease of tumor multiplicity in wild-type mice but not in Pparβ/δ-null mice. Biochemical and molecular analysis of skin and tumor samples show that these effects were due to the modulation of terminal differentiation, attenuation of inflammatory signaling, and induction of apoptosis through both PPARβ/δ-dependent and PPARβ/δ-independent mechanisms. Increased levels and activity of PPARβ/δ by nimesulide were also observed. These results are the first to demonstrate that PPARβ/δ inhibits tumorigenesis through inhibition of mitosis and promotion of cellular senescence. These studies also identify a new pro-tumorigenic role of ER stress facilitated by attenuation of senescence. Moreover, these studies support the hypothesis that combining iv

5 ligand activation of PPARβ/δ with inhibition of COX2 activity or inhibition of mitosis increases the efficacy of preventing chemically induced skin tumorigenesis as compared with either approach alone. v

6 TABLE OF CONTENTS LIST OF FIGURES...x LIST OF TABLES...xiii Chapter 1 Introduction The peroxisome proliferator-activated receptors (PPARs) PPARβ/δ expression patterns in mouse and human cancer models PPARβ/δ and the hallmarks of cancer PPARβ/δ and skin tumorigenesis PPARβ/δ regulates normal skin homeostasis Mouse skin carcinogenesis models PPARβ/δ regulates skin carcinogenesis Hypothesis Bibliography...41 Chapter 2 PPARβ/δ crosstalks with E2F and attenuates mitosis in HRAS-expressing cells Abstract Introduction Materials and methods Virus production Plasmids Cell culture Cell proliferation assay Immunofluorescence analysis Flow cytometry analysis DNA microarray analysis Gene Set Enrichment Analysis (GSEA) Quantitative western blot analysis RNA isolation and quantitative real-time PCR (qpcr) analysis Chromatin immunoprecipitation (ChIP) ChIP-re-ChIP assay Immunoprecipitation assay Luciferase reporter assay In vitro kinase assay Results Ligand activation of PPARβ/δ induces G2/M arrest causing selection against high HRAS-expressing cells Inhibition of mitosis by ligand activation of PPARβ/δ in HRAS-expressing cells Ligand activation of PPARβ/δ decreases expression of E2F target genes that regulate mitosis in HRAS-expressing keratinocytes Ligand activation of PPARβ/δ represses CDK1 and E2F1 by increasing recruitment of p107/p130 to E2F4 binding sites PPARβ/δ interacts with p107 and p Ligand activation of PPARβ/δ attenuates mitosis in vivo vi

7 2.4.7 Enhanced sensitivity to pharmacological inhibition of mitosis in HRAS-expressing cells by ligand activation of PPARβ/δ Discussion Bibliography Chapter 3 PPARβ/δ promotes HRAS-induced senescence and tumor suppression by regulating p-erk, p-akt and ER Stress Abstract Introduction Materials and methods Plasmids and Vectors Cell culture Virus production and keratinocytes infection Complete carcinogenesis bioassay In vitro malignant transformation assay Clonogenic assay with ER stress inducers Cell proliferation assays Senescence-associated β-galactosidase (SA-β-gal) assay Flow cytometry analysis Immunofluorescence analysis Protein synthesis assay Reactive Oxygen Species (ROS) detection assay RNA isolation, quantitative real-time PCR (qpcr) and RT-PCR-based detection of XBP-1 splicing Quantitative western blot analysis RAS activity assay Chromatin immunoprecipitation (ChIP) Electrophoretic mobility shift assay (EMSA) Luciferase reporter assay DNA microarray analysis Gene Set Enrichment Analysis (GSEA) Human adenoma samples Results PPARβ/δ promotes HRAS-induced senescence and suppresses malignant conversion PPARβ/δ promotes senescence by diverting from the PI3K/AKT to MEK-ERK signaling Higher p-akt decreased FOXO activity to attenuate senescence PPARβ/δ positively regulates RASGRP1 and represses ILK and PDPK1 to promote HRAS-induced senescence PPARβ/δ attenuates HRAS-induced ER stress via inhibiting p-akt-mtor activity ER stress-associated UPR inhibits HRAS-induced senescence PPARβ/δ attenuates p-akt and ER stress to promote senescence in mouse skin tumors in vivo PPARβ/δ promotes and UPR attenuates cellular senescence in human benign lesions vii

8 3.5 Discussion PPARβ/δ promotes HRAS-induced senescence by potentiating p-erk and attenuating p-akt activity PPARβ/δ dampens the negative feedback response to potentiate p-erk signaling PPARβ/δ represses ILK and PDPK1 expression Effects of ligand activation of PPARβ/δ on HRAS-induced senescence ER stress-associated UPR attenuates HRAS-induced senesce Anti-tumorigenic role of PPARβ/δ in human cancer Bibliography Chapter 4 Chemoprevention of chemically-induced skin tumorigenesis by ligand activation of peroxisome proliferator-activated receptor-β/δ (PPARβ/δ) and inhibition of cyclooxygenase 2 (COX2) Abstract Introduction Materials and Methods Two-stage chemical carcinogenesis bioassay Short-term bioassay Keratinocyte culture Caspase 3/7 activity assay Western blot analysis RNA isolation and quantitative real-time PCR (qpcr) analysis Statistical analysis Results Ligand activation of PPARβ/δ and inhibition of COX2 enhances chemoprevention of chemically-induced skin tumorigenesis Effect of GW0742 and nimesulide on terminal differentiation markers Effect of GW0742 and nimesulide on the inflammatory response Effect of GW0742 and nimesulide on apoptosis Effect of GW0742 and nimesulide on expression and function of PPARβ/δ Discussion Bibliography Chapter 5 Discussion PPARβ/δ regulates oncogenic Hras signaling in skin cancer Enhanced chemopreventive effects of combining ligand activation of PPARβ/δ with COX2 inhibition in skin carcinogenesis Future studies Characterization of endogenous PPARβ/δ ligands in HRAS-induced tumorigenesis Characterization of PPARβ/δ function in Kras-induced colon tumorigenesis Summary viii

9 5.5 Bibliography ix

10 LIST OF FIGURES Figure 1-1. Model of transcriptional regulation of PPAR....2 Figure 2-1. Ligand activation of PPARβ/δ attenuates cell proliferation by inducing G2/M arrest...76 Figure 2-2. Ligand activation of PPARβ/δ induces G2/M arrest causing selection against high HRAS-expressing cells Figure 2-3. Ligand activation of PPARβ/δ inhibits mitosis of HRAS-expressing keratinocytes...80 Figure 2-4. Ligand activation of PPARβ/δ delays mitosis entry in HRAS-expressing keratinocytes Figure 2-5. Ligand activation of PPARβ/δ increased cells with polyploidy DNA Figure 2-6. Ligand activation of PPARβ/δ decreases expression of genes that modulate mitosis in HRAS-expressing keratinocytes...85 Figure 2-7. Quantitative real-time PCR confirming decreased expression of genes that modulate mitosis in HRAS-expressing keratinocytes by ligand activation of PPARβ/δ Figure 2-8. Quantitative western blot analysis confirming decreased expression of genes that modulate mitosis in HRAS-expressing keratinocytes by ligand activation of PPARβ/δ Figure 2-9. PPARβ/δ regulates E2F target genes in HRAS-expressing keratinocytes...88 Figure PPARβ/δ regulates E2F1 in HRAS-expressing keratinocytes...89 Figure Ligand activation of PPARβ/δ inhibits expression of pro-mitotic proteins in 308 cells Figure Ligand activation of PPARβ/δ increases nuclear p107/130 protein in HRAS-expressing cells FIgure Ligand activation of PPARβ/δ represses CDK1 and E2F1 by increasing recruitment of p107/130 and represses CHEK1 by decreasing recruitment of E2F1 to respective promoters in HRAS-expressing keratinocytes Figure Effect of ligand activation of PPARβ/δ on nuclear accumulation and promoter occupancy of p130/p107 and/or E2F in 308 cells Figure PPARβ/δ binds with p107/p Figure PPARβ/δ binds with p107/p130 but not E2F4 and PPARβ/δ, p107/p130 and E2F4 form a large complex Figure Ligand activation PPARβ/δ attenuates phosphorylation of p130 by CDKs Figure Ligand activation of PPARβ/δ selects against higher HRAS-expressing chemically-induced skin tumors by inhibiting mitosis Figure Ligand activation of PPARβ/δ leads to hypersensitivity to pharmacological inhibition of mitosis in HRAS-expressing keratinocytes Figure Mechanism of PPARβ/δ mediated inhibition of mitosis in HRAS expressing cells Figure 3-1. PPARβ/δ suppresses malignant conversion x

11 Figure 3-2. PPARβ/δ promotes HRAS-induced senescence Figure 3-3. PPARβ/δ-dependent diversion from PI3K/AKT to the MEK/ERK pathway Figure 3-4. p-akt suppresses and p-erk promotes HRAS-induced senescence Figure 3-5. PPARβ/δ-dependent diversion from PI3K/AKT to the MEK/ERK pathway Figure 3-6. Higher p-akt decreased FOXO activity to attenuate senescence. 149 Figure 3-7. PPARβ/δ dampens HRAS-induced negative feedback response Figure 3-8. Rasgrp1 is a PPARβ/δ target gene Figure 3-9. PPARβ/δ represses Ilk and Pdpk Figure PPARβ/δ overexpression decreased the mrna level of both Ilk and Pdpk1 and ligand activation of PPARβ/δ did not further increase the level of Ilk or Pdpk1 in PPARβ/δ-overexpressing HaCat cells Figure PPARβ/δ positively regulates RASGRP1 and represses ILK to promote senescence Figure PPARβ/δ attenuates HRAS-induced ER stress Figure PPARβ/δ attenuates HRAS-induced ER stress without affecting apoptosis Figure PPARβ/δ attenuates HRAS-induced ER stress via inhibiting p-akt activity Figure PPARβ/δ attenuates HRAS-induced ER stress via inhibiting p-akt-mtor activity Figure Decreased XBP-1 increased HRAS-induced cellular senescence Figure Decreased ATF4 increased HRAS-induced cellular senescence. 162 Figure Increased ER stress-associated UPR by thapsigargin attenuates HRAS-induced senescence Figure Increased ER stress-associated UPR attenuates HRAS-induced senescence through a p-akt dependent mechanism Figure A brief increase of ER stress-associated UPR is sufficient to attenuate HRAS-induced senescence Figure UPR increases p-akt partially through cell-surface BiP Figure A brief increase of UPR leads to malignant conversion in vitro Figure PPARβ/δ promotes cellular senescence in mouse skin tumors in vivo Figure PPARβ/δ attenuates ER stress in mouse skin tumors in vivo Figure ER stress and UPR negatively correlates with cellular senescence in mouse skin tumors in vivo Figure PPARβ/δ attenuates p-akt and ER stress to promote senescence in vivo Figure PPARβ/δ promotes and UPR attenuates cellular senescence in human benign lesions Figure A model of regulation of Hras-induced senescence by PPARβ/δ..185 Figure 4-1. Chemoprevention of chemically-induced skin tumorigenesis by combining ligand activation of PPARβ/δ and inhibition of COX xi

12 Figure 4-2. Skin tumor size following ligand activation of PPARβ/δ and inhibition of COX Figure 4-3. Distribution of keratoacanthomas and squamous cell carcinomas following ligand activation of PPARβ/δ and inhibition of COX Figure 4-4. Histopathology of skin tumors following ligand activation of PPARβ/δ and inhibition of COX Figure 4-5. Expression of differentiation markers in skin following ligand activation of PPARβ/δ and inhibition of COX Figure 4-6. Effect of ligand activation of PPARβ/δ and/or inhibition of COX2 on Il6 and Tnfα mrna in skin tumors and skin Figure 4-7. Effect of ligand activation of PPARβ/δ and/or inhibition of COX2 on apoptotic signaling Figure 4-8. Effect of ligand activation of PPARβ/δ and/or inhibition of COX2 on expression and activity of PPARβ/δ xii

13 LIST OF TABLES Table 1. Key aspects of hallmarks of cancer that PPARβ/δ regulates...29 xiii

14 Chapter 1 Introduction In the first chapter, I will first introduce the basic concepts of the peroxisome proliferator-activated receptors (PPARs). Next, I will discuss the relationship between PPARβ/δ and cancer. In this section, I will first talk about the expression patterns of PPARβ/δ in different human and mouse cancer models. Next,I will elaborate on the role of PPARβ/δ in regulating different hallmarks of cancer, a concept first introduced by Robert Weinberg and expanded by Stephen Elledge (Hanahan and Weinberg, 2000; Hanahan and Weinberg, 2011; Kroemer and Pouyssegur, 2008; Luo et al., 2009). Finally, I will focus on the regulatory functions of PPARβ/δ in non-melanoma skin tumorigenesis The peroxisome proliferator-activated receptors (PPARs) The peroxisome proliferator-activated receptors (PPARs) belong to the type II class of nuclear receptor super family that function as transcription factors regulating the expression of target genes. PPARs play essential roles in the regulation of adipogenesis, lipid metabolism, cell proliferation/apoptosis, cell differentiation, inflammatory responses and carcinogenesis (Berger and Moller, 2002; Feige et al., 2006). The PPARs consist of three distinct isoforms, PPARα, PPARβ/δ (also refered to as PPARβ or PPARδ) and PPARγ. PPARs are found in both cytosol and nucleus, often forming complexes with co-repressors in the absence of ligands (Shi et al., 2002). On ligand activation, they undergo conformational changes followed by the dissociation of co-repressors, recruitment of co-activators and heterodimerization with the retinoid X receptor-α (RXRα) (Yu 1

15 and Reddy, 2007). The heterodimer then binds to the peroxisome proliferator response element (PPRE) in the target genes and turns on the transcription process (Yu and Reddy, 2007) (Figure 1-1). In addition, there is also evidence showing that PPARβ/δ can repress certain genes expression by binding to PPREs in the presence of co-repressors (Shi et al., 2002). Interestingly, PPARβ/δ has been shown to be able to repress PPARα and PPARγ target gene transactivation by competing for the PPREs (Shi et al., 2002). Recent evidences also revealed that the PPARs can interact with p65 subunit of NF-κB, AP1 and STAT3 and interfere with thesesignaling pathways (Konstantinopoulos et al., 2007; Wang et al., 2005; Wang et al., 2002). Figure 1-1. Model of transcriptional regulation of PPAR. In response to ligand activation, peroxisome proliferator-activated receptors (PPARs) heterodimerize with retinoid X receptor (RXR), dissociate with corepressor complex and recruit transcriptional machinery, including RNA polymerase and co-activators with histone acetyl transferase activity, leading to remodelling of chromatin and increased transcription. 2

16 Of the three isoforms, the biological functions of PPARα and PPARγ are more clearly characterized. PPARα is expressed in liver, kidney, heart, muscle, and adipose tissue, where it is known to stimulate lipid catabolism by activating genes involved in peroxisomal β-oxidation (Kersten et al., 1999; Mukherjee et al., 1994), PPARγ is expressed in adipose tissue and regulates adipogenesis (Feige et al., 2006). PPARβ/δ is expressed with higher levels detected in liver, adipose tissue, intestine and skin (Burdick et al., 2006; Girroir et al., 2008a). There is good evidence suggesting that PPARβ/δ plays an important role in regulating metabolism. For example, ligand activation of PPARβ/δ has been shown to raise high density lipoprotein (HDL)-cholesterol and decrease low density lipoprotein (LDL)-cholesterol in two different obese animal models (Leibowitz et al., 2000; Oliver et al., 2001). Furthermore, ligand activation of PPARβ/δ also can prevent obesity and increase insulin sensitivity by facilitating fatty acid β-oxidation in skeletal muscle and adipose tissue (Tanaka et al., 2003; Wang et al., 2003), suppressing glucose output and increasing glucose disposal in the liver (Lee et al., 2006) and mediating macrophage alternative activation (Kang et al., 2008; Odegaard et al., 2008), which is believed to improve insulin sensitivity. A recent paper also showed that the activation of PPARβ/δ together with exercise training can increase running endurance by increasing oxidative fibers (Narkar et al., 2008). Together these reports suggest that PPARβ/δ agonists have therapeutical values for treating metabolic disorders. 3

17 1.2. PPARβ/δ expression patterns in mouse and human cancer models There is conflicting literature regarding the expression level of PPARβ/δ in human and mouse cancers. A number of papes report higher levels of PPARβ/δ expression in colon cancers compared to normal tissue (Delage et al., 2007; Gupta et al., 2000; He et al., 1999; Roy et al., 2001; Takayama et al., 2006; Yoshinaga et al., 2009). The increased expression of PPARβ/δ was attributed to the enhanced APC-β-catenin-TCF-mediated transcription upregulation of PPARβ/δ, which was reported to be a target gene of this pathway (He et al., 1999). It was further hypothesized that increased PPARβ/δ expression promoted colon tumorigenesis in these reports. Other studies also reported increased expression of PPARβ/δ in cancer tissues as compared to normal tissues in ovarian carcinomas, squamous cell carcinomas and breast cancers (Davidson et al., 2009; Glazer et al., 2008; Jaeckel et al., 2001; Nijsten et al., 2005). However, most of the conclusions were based solely upon immunohistochemistry of PPARβ/δ or just quantification of mrna level of PPARβ/δ. The limitation of immunohistochemistry arose in part from the unspecificity of commercially available anti-pparβ/δ antibodies. In addition, the mrna level of PPARβ/δ does not necessarily correlate with that of protein level since PPARβ/δ is known to be regulated post-translationally (Genini and Catapano, 2007). On the other hand, a number of studies also reported decreased or unchanged level of PPARβ/δ in transformed colons compared to normal tissue and provided evidence arguing against the hypothesis that PPARβ/δ is a downstream target of APC-β-catenin-TCF pathway (Baek et al., 2004; Chen et al., 2004; Foreman et al., 4

18 2011; Hao et al., 2005; Modica et al., 2010; Notterman et al., 2001; Reed et al., 2004; Yang et al., 2011a). Recently, a report on the journal of clinical cancer research provided the most convincing and elegant report on the expression and functional of PPARβ/δ in human colorectal lesions to date (Yang et al., 2011a). In this report, it was shown that PPARβ/δ expression was higher in some primary cancers when compared to normal colon tissue but its expression was reduced in lymph node metastasis (Yang et al., 2011a), which was also consistent with another early report (Yang et al., 2010). In addition, higher expression of PPARβ/δ was correlated with decreased expression of cell proliferation marker Ki-67 (Yang et al., 2011a). More importantly, patients of rectal cancer with lower expression of PPARβ/δ were four times more likely to die of colorectal cancer than patients with relatively higher expression of PPARβ/δ. Studies in our lab using qrt-pcr and quantitative western blot analysis also showed that both mrna and protein level of PPARβ/δ was lower in human malignant colon adenocarcinomas compared to normal tissue, which is also consistent with a recent report by Modica et al (Modica et al., 2010). Further, stable over-expression of PPARβ/δ in human colon cancer cell lines enhanced ligand activation of PPARβ/δ and inhibition of clonogenicity in HT29 colon cancer cells (Foreman et al., 2011). These findings provided strong evidence that PPARβ/δ may attenuate colon tumorigenesis rather than promote it PPARβ/δ and the hallmarks of cancer The original hallmarks of cancer, first proposed in a seminal paper in Cell in 2000, 5

19 include sustaining proliferative signaling, evading growth suppressors, resisting cell death, enabling replicative immortality, inducing angiogenesis, and activating invasion and metastasis (Hanahan and Weinberg, 2000). These traits are distinct characteristics of cancer cells that distinguish them from normal cells. Later on, evasion of immune surveillance was added as an additional aspect of cancer hallmark (Kroemer and Pouyssegur, 2008). In 2009, a set of additional hallmarks that depict the stress phenotypes of cancer cells, including metabolic stress, proteotoxic stress, mitotic stress, oxidative stress and DNA damage stress, were added (Luo et al., 2009). Furthermore, tumor-promoting inflammation was also considered to be an important enabling characteristic for tumor growth (Hanahan and Weinberg, 2011). Together, these thirteen hallmarks of cancer designate the distinct traits of cancer cells and also shape the framework for finding new targets for cancer therapeutic interventions. (Hanahan and Weinberg, 2000). PPARβ/δ can regulate tumorigenesis by affecting virtually every hallmark of cancer. In the following sections, I will elaborate on each hallmark in detail and review some of the literature linking PPARβ/δ to that specific hallmark. More importantly, I will focus on the mechanistic regulation of each hallmark by PPARβ/δ, rather than just showing expression patterns of PPARβ/δ in different cancers or a mere involvement of PPARβ/δ in each hallmark. Sustained angiogenesis Due to the rapid growth of tumor tissues, a continual supply of oxygen and nutrients to the tumor tissue by blood vessels is essential for both maintaining the primary tumor and metastasis to a secondary site. Therefore, tumors develop 6

20 angiogenic ability by upregulating angiogenesis inducers and countervailing inhibitors. These include but are not limited to increased expression of vascular endothelial growth factor (VEGF) and acetic and basic fibroblast growth factor (FGF 1/2) and down-regulation of angiogenesis inhibitors such as thrombospondin-1 (Hanahan and Weinberg, 2000). Two studies linked PPARβ/δ to angiogenesis (Abdollahi et al., 2007; Muller-Brusselbach et al., 2007). In the absence of PPARβ/δ, an abundance of morphologically immature and hyperplastic microvessels were found surrounding tumors, causing a diminished blood flow and subsequent impaired tumor growth. The proliferating nature of endothelial cells in the absence of PPARβ/δ was partially caused by decreased expression of p57 Kip2, a cell cycle inhibitor believed to be a PPARβ/δ target gene (Muller-Brusselbach et al., 2007). In addition, decreased expression of thrombospondin-1 and its receptor CD36 were also found in Pparβ/δ-null cells even though the mechanism of their regulation by PPARβ/δ remains unknown (Muller-Brusselbach et al., 2007). Furthermore, ligand activation of PPARβ/δ can either increase VEGF or decrease VEGF receptor expression in cancer and endothelial cells, subsequently increasing or decreasing proliferation of endothelial cells, potentially causing increased or decreased angiogenesis (Fauconnet et al., 2002; Meissner et al., 2010). Therefore, there is still no consensus on the role of PPARβ/δ in angiogenesis and whether PPARβ/δ is promoting or inhibiting tumorigenesis through its effects on angiogenesis remains unclear. 7

21 Tissue invasion and metastasis Cancer cells can break away from their site or organ of origin to invade surrounding tissues and spread to distant body parts. Metastasis is complex process that involves multiple stages and cooperation of a plethora of gene expression alterations. For example, E-cadherin, expressed on the cell surface of epithelial cells mediates cell-cell junction and is widely believed to be suppressors of invasion and metastasis and its expression is usually decreased in order for cancer cells to invade surrounding tissues. In addition, cancer cells exhibit an epithelial to mesenchymal transition (EMT) during tissue invasion and metastasis (Chaffer and Weinberg, 2011). Currently, the evidence suggesting a role of PPARβ/δ in mediating metastasis is lacking. However, several observations may imply some potential regulatory roles of PPARβ/δ in metastasis. First, Angiopoietin-like 4 (Angptl4), one of the most well-characterized target gene of PPARβ/δ, has been shown to have either pro or anti-metastasis functions (Galaup et al., 2006; Padua et al., 2008). Tumor cell-derived ANGPLT4 has been shown to disrupt vascular endothelial cell-cell junctions, increases the permeability of lung capillaries, and facilitates the trans-endothelial passage of breast tumor cells into lungs (Padua et al., 2008). In contrast, ANGPTL4 can also prevent metastasis through inhibition of vascular permeability and tumor cell motility and invasiveness (Galaup et al., 2006). So it is likely that PPARβ/δ may mediate metastasis through an ANGPTL4-dependent mechanism. However, it should be noted that PPARβ/δ expression does not necessarily correlate with its activity nor with ANGPLT4 expression since the 8

22 presence or level of endogenous ligand for PPARβ/δ may vary in different tumor types. Secondly, expression of several members of matrix metallopeptidase (MMP) including MMP3, MMP10 and MMP13 was higher in Pparβ/δ-null keratinocytes (unpublished data). MMPs have been implicated in tumor metastasis by their abilities to degrade extra-cellular matrix. So it is equally likely that PPARβ/δ may inhibit metastasis by repressing the expression of MMPs. Indeed, a recent paper showed that PPARβ/δ agonists potently inhibited TGFalpha-induced MMP9 expression in human keratinocytes through an AP-1 dependent mechanism (Meissner et al., 2011). However, more extensive in vivo studies are needed to shed light on the potential roles of PPARβ/δ in mediating metastasis using both mouse and human models Evading apoptosis Cancer cells experience various physiological stresses that may trigger apoptosis and they have evolved multiple strategies to avoid it. The most common one is the loss of p53 tumor suppressor function. Loss of p53 eliminates the critical damage sensor from the apoptosis-inducing machinery (Vazquez et al., 2008). Alternatively, tumors may circumvent apoptosis by increasing expression of anti-apoptotic regulators (Bcl-2, Bcl-xL) or of survival signals (Igf1/2), by down-regulating pro-apoptotic factors (Bax, Bim, Puma), or by impairing the extrinsic ligand-induced death pathway (Hanahan and Weinberg, 2011). The role of PPARβ/δ in modulating apoptosis remains controversial due to disparities in literature. One study suggesting anti-apoptotic role of PPARβ/δ 9

23 showed that ligand activation PPARβ/δ increased the expression of integrin-linked kinase (ILK) and 3-phosphoinositide-dependent-protein kinase 1 (PDPK1) and decreased the expression of PTEN, leading to increased phosphorylation of AKT and subsequent increased anti-apoptotic and pro-survival capacity (Di-Poi et al., 2002). However, following studies in our lab and others showed no increased expression of ILK and PDPK1 after ligand activation of PPARβ/δ. In contrast, increased expression of ILK and PDPK1 was found in the absence of PPARβ/δ (Burdick et al., 2007; Marin et al., 2006; Yang et al., 2010). Therefore the proposed anti-apoptotic function of PPARβ/δ through increased AKT signaling was not found in these studies. A similar study showed that ligand activation of PPARβ/δ increased the expression of vascular endothelial growth factor (VEGF) through a PPARβ/δ-dependent mechanism, causing increased phospho-akt level, which in turn promoted cell survival by blocking apoptosis (Wang et al., 2006). However, results in our lab showed that ligand activation of PPARβ/δ did not increase the expression of VEGF nor increase the phosphorylation of AKT in human cancer cell lines (Hollingshead et al., 2007). In addition, it was reported that increased PPARβ/δ expression and/or ligand activation of PPARβ/δ antagonized the pro-apoptotic effects of PPARγ by counteracting the decreased survivin expression and increased caspase-3 activity mediated by PPARγ activation in colorectal cell lines (Wang et al., 2011). The authors made this conclusion largely based on the comparison of PPARβ/δ expression in two different colon cancer cells lines HCT116 and LS174T (Wang et al., 2011). While LS174T cell line was more sensitive to PPARγ 10

24 activation-induced apoptosis, HCT116 was refractory to this stimulus (Wang et al., 2011). The finding that PPARβ/δ expression was much higher in HCT116 cell line compared to LS174T cell line suggested that PPARβ/δ conferred this resistance (Wang et al., 2011). However, we and other labs showed that PPARβ/δ expression level was similar between these two cell lines but in contrast a higher PPARγ expression was found in LS174T cell lines (Foreman et al., 2009; He et al., 1999). This suggests that the difference in PPARγ expression rather than that of PPARβ/δ was responsible for the differential resistance to PPARγ-induced apoptosis in these two cell lines. Further, it was shown that nonsteroidal anti-inflammatory drugs (NSAID) increased apoptosis by decreasing expression of PPARβ/δ and increasing ε, which allows BAD to function as a pro-apoptotic protein in human colon cancer cell lines. (Liou et al., 2007; Wu and Liou, 2009). However, result from our lab showed that NSAIDs does not decrease PPARβ/δ expression and neither NSAIDs or ligand activation of PPARβ/δ changed the level of ε (Foreman et al., 2011). Due to the disparate results in various studies, there is no consensus on the function of PPARβ/δ in modulating apoptosis and extensive in vivo studies using mouse and human models are needed to address this question in the future. Self-sufficiency in growth signal Compared to normal cells, cancer cells can acquire the ability to proliferate independent of exogenous growth signals and succeed in sustaining proliferative signaling through various mechanisms (Hanahan and Weinberg, 2011). For 11

25 example, cancer cell can bypass the need for ligand activation of cell-surface receptors by constitutive activation of components downstream of these growth receptors (Hanahan and Weinberg, 2011). Alternatively, they can both produce growth factors and upregulate cognate receptors themselves, resulting in autocrine stimulation of cell growth (Hanahan and Weinberg, 2011). A third possibility involves the disruption of negative feedback loops that originally exists in normal cells to turn off the proliferative signaling when necessary (Hanahan and Weinberg, 2011). Combined, these mechanisms confer cancer cells a phenotype characterized by self-sufficiency in growth signals. PPARβ/δ regulates cancer cell proliferation through multiple mechanisms. PPARβ/δ could mediate downstream kinase pathways independent of cell surface receptors. For example, it was reported that the PPARβ/δ upregulated ubiquitin C expression to attenuate TPA-induced skin hyperplasia in wild-type mice (Kim et al., 2004a). PPARβ/δ promoted ubiquitin-dependent proteosome-mediated protein turnover of PKCα in mouse skin in response to TPA, resulting in a lower PKCα level, decreased Raf1, and MAPK/extracellular signal-regulated kinase activities and subsequent decreased cell proliferation (Kim et al., 2005a). PPARβ/δ also affected growth factor-receptor interactions in an autocrine fashion. Dermal fibroblast PPARβ/δ inhibited the mitotic activity of keratinocytes via inhibition of the IL-1 signaling pathway by transcriptional upregulation of secreted IL-1 receptor antagonist, causing an autocrine decrease in IL-1 signaling pathways and consequently decreases production of secreted mitogenic factors by the fibroblasts (Chong et al., 2009). Thus, the present 12

26 literature suggestes that PPARβ/δ exerts anti-tumorigenic functions in skin carcinogenesis in part through a mechanism involving putting a restraint on growth-factor activated downstream pathways. Insensitivity to anti-growth signals In addition to the ability to sustain proliferative signaling, cancer cells can also circumvent powerful programs that negatively regulate pro-proliferation signaling, most of which are elicited by tumor suppressors genes. Indeed, mutations in p53, RB, APC, NF1, PTEN and many other tumor suppressors have been detected in a number of human cancers (Sherr, 2004). The literature indicating that PPARβ/δ directly regulates tumor suppressor expression is lacking. In one example, PPARβ/δ was shown to facilitate CRE-binding protein (CREB)-mediated induction of p21/p27 gene expression through a PPRE-dependent mechanism (Sue et al., 2009). Another example, which is also mentioned earlier, showed that PPARβ/δ-mediated downregulation of PTEN in primary keratinocytes partially accounts for the increased anti-apoptotic and pro-survival capacity endowed by ligand activation of PPARβ/δ (Di-Poi et al., 2002). However, as indicated above, we and other labs have not been able to repeat this finding (Burdick et al., 2007; Marin et al., 2006; Yang et al., 2010). Recently, it has been shown that in smooth muscle cells, ligand-activated PPARβ/δ increased PTEN expression and subsequently decreased PI3K-AKT signaling and this led to an decrease in ROS levels (Kim et al., 2011a). Therefore, under different cellular contexts, PPARβ/δ may differentially regulate PTEN 13

27 expression. As described earlier, PPARβ/δ was first thought to be a target gene in the APC-β-CATENIN-TCF pathway to promote colon tumorigenesis in the absence of functional tumor suppressor APC (He et al., 1999). However, subsequent studies argued against this hypothesis (Baek et al., 2004; Chen et al., 2004; Foreman et al., 2011; Hao et al., 2005; Modica et al., 2010; Notterman et al., 2001; Reed et al., 2004; Yang et al., 2011a). Given the global transcriptional regulation capacity of PPARβ/δ, it would not be surprising that more tumor suppressors gene will be found that are directly or indirectly regulated by PPARβ/δ. In addition, attention should be given to those genes that are not traditionally classified as canonical tumor suppressor genes but may serve tumor suppressing functions through indirect mechanisms. Limitless replicative potential In contrast to normal cells, cancer cells acquire unlimited replicative potential in order to generate macroscopic tumors. This is accomplished mainly through the ability to evade replicative senescence, a trait that is tightly regulated by the telemeres protecting the ends of chromosomes (Blasco, 2005). Further, it is also known that eroded telemeres generate a persistent DNA damage response that elicits and maintains senescence response partially through induction of p53 tumor suppressor (Bartkova et al., 2006; Rodier and Campisi, 2011). Cancer cells evade DNA damage response-induced senescence through mutations in p53, ATM and other tumor suppressor genes (Bartkova et al., 2006; Rodier and 14

28 Campisi, 2011). In addition to replicative senescence, introduction of oncogenes into primary cells can also triggers cellular senescence response, a phenomenon first observed in mouse embryonic fibroblasts expressing oncogenic Hras (Serrano et al., 1997). Since this observation, oncogene-induced senescence has been observed in response to a number of mutations in other genes including Kras, Braf, Pten and Nf1(Collado and Serrano, 2010). It is widely believed that oncogene-induced senescence serves as a self-defense mechanism to suppress tumor development by preventing the progression of benign lesions to malignancies in the absence of additional cooperating mutations (Courtois-Cox et al., 2008). Secondary mutations targeting key tumor suppressors such as RB, p53 and p16 can allow pre-malignant lesions to bypass senescence checkpoint and eventually lead to malignancy conversion (Courtois-Cox et al., 2008). Three closely related papers link PPARβ/δ with DNA damage response-induced senescence response (Ham et al., 2012; Kim et al., 2011a; Kim et al., 2011b). In smooth muscle cells, ligand activation of PPARβ/δ decreased the generation of reactive oxygen species (ROS) and suppressed angiotensin II-induced senescence (Kim et al., 2011a; Kim et al., 2011b). Ligand-activated PPARβ/δ decreased the generation of ROS through two not mutually exclusive mechanisms. First, ligand-activated PPARβ/δ increased PTEN expression and subsequently decreased PI3K-AKT signaling and this led to an decrease in ROS levels (Kim et al., 2011a). In addition, ligand activation of PPARβ/δ by GW upregulated the expression of antioxidant genes including glutathione peroxidase 1, thioredoxin 1, manganese superoxide dismutase and heme oxygenase 1, 15

29 which accounted for the decreased ROS (Kim et al., 2011b). However, whether this is due to a PPARβ/δ-dependent mechanism or it is just ligand specific effect is not known. Furthermore, it was also not known whether PPARβ/δ directly regulated these antioxidant genes or through cross talk with other transcription factors. In human keratinocytes, ligand activation of PPARβ/δ decreased the generation of reactive oxygen species (ROS) and suppressed UVB-induced senescence, a phenomenon also mediated through ligand-dependent attenuation of PI3K-AKT pathway (Ham et al., 2012). It is worth pointing out that none of the three papers included PPARβ/δ loss-of-function controls, therefore the possibility that all these effects were caused by off target effects of GW alone cannot be ruled out. With the emerging evidence suggesting that PPARβ/δ regulates DNA damage response-induced cellular senescence response, it is tempting to hypothesize that PPARβ/δ may also regulate cellular senescence response triggered by other stimuli, such as telemere shortening or aberrant oncogenic signaling. In addition to cell culture models, mouse and human models should be utilized to further test this hypothesis. DNA damage stress and oxidative stress Cancer cells, especially solid tumor cells experience stages of extreme genomic instability that result in the accumulation of point mutations, deletions and even chromosomal rearrangements (Hartwell and Kastan, 1994). This level of instability is due partially to constitutive level of endogenous DNA damage that 16

30 activates the DNA damage response outlined in the previous section (Bartkova et al., 2005). Factors that can cause this DNA damage in early stage of tumorigenesis include the double-strand breaks at telomeric ends due to the shortening of telomeres in the absence of sufficient telomerase activity (Luo et al., 2009). Additionally, oncogene activation in premalignant tumors could also increase double-strand breaks and genomic instability (Halazonetis et al., 2008). Finally, mutation of genes involved in either DNA repair programs (BRCA1, BRCA2) or the DNA damage response pathways (such as ATM and p53) has been known to increase the DNA damage (Harper and Elledge, 2007). Coupled with the ability to evade DNA damage response-induced senescence, cancer cells can continue to proliferate in the presence of DNA damage, which in turn can increase the chance of secondary mutations, further facilitating the tumorigenesis process. In addition to the cell autonomous mechanisms that lead to an accumulation of DNA damage, foreign carcinogens also can cause DNA damage, initiate and promote tumorigenesis. In fact, exposure to chemical carcinogens contributes significantly to both cancer and toxicities (Wogan et al., 2004). One of the central regulators in metabolizing xenobiotics, which include a variety of carcinogens, is the aryl hydrocarbon receptor (AHR). The aryl hydrocarbon receptor (AhR) mediates increased expression of phase I and II xenobiotic metabolizing enzymes in response to exposure to chemical carcinogens including polycyclic aromatic hydrocarbons (PAH) (Rowlands and Gustafsson, 1997). The balance between bioactivation and detoxification determines the extent of DNA damage following 17

31 exposure to carcinogens. Thus far there is no literature available that connects PPARβ/δ to telomere shortening or double strand breaks-induced DNA damage response in cancer cells. However, as previously described, ligand activation of PPARβ/δ could decrease the generate of ROS in smooth muscle cells and keratinocytes (Ham et al., 2012; Kim et al., 2011a; Kim et al., 2011b). Therefore it is possible that PPARβ/δ may also regulate DNA damage response in cancer cells through a similar mechanism. However it is worth pointing out that the functional consequences of decreasing DNA damage in cancer cells are controversial. On one hand, decreasing DNA damage stress could exert pro-survival effects by relieving cancer cells of exacerbated stress (Bartkova et al., 2005). On the hand other, it may also attenuate tumorigenesis by decreasing the possibility of creating secondary mutations that are necessary for late stages of tumor growth (Jackson and Loeb, 2001). Therefore, accurate quantification and functional analysis of effects of ligand activation of PPARβ/δ on ROS generation and establishing a direct correlation between ROS and tumor growth in mouse models is needed to fully elucidate this problem. It has been found in our lab that PPARβ/δ can alter AhR-dependent signaling by modulating cytochrome P450 (CYP) expression. In the absence of PPARβ/δ expression, increased expression of CYP1B1 and CYP1A1 and some phase II enzymes does not occur after application of PAHs (unpublished data). Since there is a balance between bio-activation (phase I) and detoxification (phase II) of carcinogens that is mediated by AhR-dependent pathways, this suggests that 18

32 PPARβ/δ could significantly alter this balance and thus influence chemically- induced carcinogenesis accordingly. The mechanism by which PPARβ/δ regulates AhR signaling is still undergoing investigation. Mitotic stress A subset of tumors undergo chromosome mis-segregation during mitosis, a phenomenon referred to as chromosome instability (Komarova et al., 2002). Chromosome instability could result from defects in a variety of pathways involved in mitosis that include mitotic proteins that execute chromosome segregation and proteins involved in spindle assembly checkpoint. In addition, mutations in genes regulating mitosis have been found in various human cancers (Perez de Castro et al., 2007). All of this evidence suggests that cancer cells are strongly addicted to the enhanced mitotic machinery and as a result they undergo severe mitotic stress. The only evidence suggesting a role of PPARβ/δ in regulating mitosis thus far was the finding that ligand activation of PPARβ/δ induces a G2/M block in cancer cells with an activating Hras mutation (Bility et al., 2010). It is likely that PPARβ/δ induces G2/M block by blocking mitosis and functional study is needed to prove this hypothesis. Proteotoxic stress Tumors exhibit proteotoxic stress evidenced by a constitutive display of heat shock response and enhanced ER stress. Elevated hear shock response is 19

33 thought to be caused by aneuploidy often observed in cancer cells (Ganem et al., 2007; Torres et al., 2008). The rationale for this theory is that aneuploidy often leads to increased or decreased levels of growth and survival signals and eventually result in an imbalance in the stoichiometry of protein complex subunits (Papp et al., 2003). ER stress, on the other hand, can be triggered by a variety of stimuli, including but not limited to deprivation of glucose, changes in redox potential, perturbations of calcium level in the cells, and direct oncogenic signaling (Lin et al., 2008). ER stress can in turn elicit a similar response called unfolded protein response to facilitate proper protein folding (Lin et al., 2008). In a sense, both heat shock response and unfolded protein response can be considered as the same response since they both converge on regulating protein-folding homeostasis. Originally, both heat shock response and unfolded protein response were considered to serve pro-survival roles in cancer cells (Tsai and Weissman, 2010). However, when the level of ER/proteotoxic stress exceeds a certain threshold that can no longer be tolerated by cells, an apoptosis program will be activated (Tsai and Weissman, 2010). Therefore, under different cellular contexts, ER/proteotoxic stress can promote or inhibit tumor growth (Tsai and Weissman, 2010). Even though there is no literature at present that implies that PPARβ/δ regulates heat shock response, unpublished preliminary data in our lab showed that Pparβ/δ-null keratinocytes are more sensitive to heat shock-induced cell death as compared to wild-type cells. This is partially due in part to decreased induction of heat shock protein 70 in Pparβ/δ-null keratinocytes in response to heat shock as 20

34 compared to wild-type keratinocytes. Additionally, due to the regulation of ubiquitin C gene expression by PPARβ/δ (Kim et al., 2004a), a decrease in free ubiquitin pool in Pparβ/δ-null keratinocytes potentially may also underlie the exacerbated heat shock stress observed in Pparβ/δ-null keratinocytes. However, more functional experiments are needed to prove this hypothesis. Further, whether this differential regulation of heat shock response by PPARβ/δ in normal keratinocytes could translate into a functional consequence in cancer cells remains to be determined. Metabolic stress Normal cells utilize mitochondrial oxidative phosphorylation to generate ATP (Kalckar, 1974). Most cancer cells, on the other hand, are found to predominantly produce energy by the less efficient method of glycolysis and secrete a large amount of lactic acid, even under high oxygen conditions (Warburg, 1956), This effect is referred to as Walburg effects or aerobic glycolysis. Tumor cells exhibit increased glucose uptake and increased rates of glycolysis (DeBerardinis et al., 2008; DeBerardinis et al., 2007). This diversion to glycolysis for energy production provides tumor cells with several advantages. First, it prepares tumors cells for the potential low oxygen environment often observed in late stage of tumorigenesis (DeBerardinis et al., 2008; Kroemer and Pouyssegur, 2008). Secondly, glycolysis could decrease ROS generation by both the decreased use of mitochondrial oxidative phosphorylation and by the generation of NADPH, the major reducing agents, to synthesize anti-oxidant glutathione through the pentose 21

35 phosphatase pathway (DeBerardinis et al., 2008; Kroemer and Pouyssegur, 2008). Thirdly, increased glycolysis also can divert resources toward biosynthesis such as fatty acid synthesis (DeBerardinis et al., 2008; Kroemer and Pouyssegur, 2008). Lastly, the acidification of the surrounding microenvironment by glycolysis could promote tumor invasion and suppresses immune surveillance (Luo et al., 2009). As mentioned earlier, PPARβ/δ plays important roles in regulating metabolism. In the liver, ligand activation of PPARβ/δ suppresses hepatic glucose output, increases glucose disposal by increasing glucose flux through the pentose phosphate pathway and enhancing fatty acid synthesis (Lee et al., 2006), both of which are also occurring in most cancer cells. Similarly, ligand activation of PPARβ/δ also increased several fatty acid synthesis gene expression (unpublished data) and stimulated lipid accumulation (Schmuth et al., 2004) in mouse primary keratinocytes. However, conflicting results also demonstrated that PPARβ/δ induces insulin-induced gene-1 and suppresses hepatic lipogenesis in obese diabetic mice (Qin et al., 2008). In skeletal muscle and adipose tissue, ligand activation of PPARβ/δ was shown to prevent obesity and increase insulin sensitivity by facilitating fatty acid β-oxidation (Kleiner et al., 2009; Tanaka et al., 2003; Wang et al., 2003), thus reflecting distinct regulation of metabolism in different tissues. Given the large amount of evidence suggesting pivotal roles of PPARβ/δ in regulating metabolism, it is surprising that no study has ever examined the hypothesis that PPARβ/δ regulate metabolism to affect tumorigenesis in vitro or in 22

36 vivo. However, it is tempting to hypothesize that PPARβ/δ may favor liver tumorigenesis by increasing glucose flux through the pentose phosphate pathway and facilitating fatty acid synthesis. On the other hand, given the important pro-tumorigenic functions of chronic inflammation that will be fully discussed below, it is equally possible that PPARβ/δ may also systematically attenuates tumorigenesis through facilitating the maintenance of a lean phenotype and increased insulin sensitivity and thus decreasing the chronic inflammation normally associated with obesity, Future studies are needed to prove these hypothesizes. Immune system and cancer The relationship between the immune system and cancer is complex. On one hand, chronic inflammation has long been known to stimulate tumor growth and progression, mainly through secreting pro-proliferative cytokines such as IL-1, IL-6 and TNF-α (Grivennikov et al., 2010). However, cancer cells also represent an altered self and display foreign antigens that can be recognized by immune system, therefore growing tumors are also subject to immunosurveillance by activated T and NK cells (Grivennikov et al., 2010). The balance between the pro-tumorigenic effects of inflammation and anti-tumorigenic effects of immunosurveilance may underlie the long period of tumor dormancy (Koebel et al., 2007). Moreover, tumor cells often edit their tumor antigens in late stage of tumor growth to escape immunosurveillance, thus tilting the balance to favor tumor growth (Koebel et al., 2007). 23

37 There is a large body of evidence suggesting anti-inflammatory effects of ligand activation of PPARβ/δ in different models. At least two mechanisms account for the inhibition of pro-inflammatory signaling by ligand activation of PPARβ/δ. First, ligand activation of PPARβ/δ can interfere with NF-κB signaling by sequestering p65 and thus subsequently inhibiting expression of pro-inflammatory cytokines such as TNF-α, IL1β and IL6 and chemokines such as monocyte chemotactic protein 1 (MCP1) and CXCL7 (Barish et al., 2008; Kilgore and Billin, 2008a; Piqueras et al., 2009; Planavila et al., 2005; Rival et al., 2002; Shan et al., 2008a; Shan et al., 2008b; Zhu et al., 2010). However, It is worth pointing out that some of the anti-inflammatory effects of ligands are PPARβ/δ independent (Zhu et al., 2010). In addition, PPARβ/δ ligand GW0742 was shown to be an uncompetitive inhibitor of MPO activity independent of PPARβ/δ (Kim et al., 2006a). Secondly, PPARβ/δ can also mediate activation of alternatively-activated anti-inflammatory M2 phenotype of macrophages (Kang et al., 2008; Odegaard et al., 2008). In spite of convincing evidence endowing PPARβ/δ with anti-inflammatory functions, whether activating PPARβ/δ could prevent tumorigenesis by inhibiting inflammation has yet to be determined. At present, whether PPARβ/δ modulate NK, NKT or cytotoxic CD8 + T cells function to affect immunosurveillance remains unknown. However, three recent papers described the novel role of PPARβ/δ in modulating experimental autoimmune encephalomyelitis (EAE) (Dunn et al., 2010; Kanakasabai et al., 2010a; Kanakasabai et al., 2010b). While ligand activation of PPARβ/δ with GW0742 ameliorated experimental autoimmune encephalomyelitis (EAE) by 24

38 blocking interferon (IFN)-gamma and interleukin (IL)-17 production by T helper type 1 (Th1) and Th17 cells (Kanakasabai et al., 2010a), Pparβ/δ-null mice showed prolonged EAE with augmented neural antigen-specific Th1/Th17 responses and elevated expression of IFNγ and IL-17 (Dunn et al., 2010; Kanakasabai et al., 2010b). Given the important role of T helper type 1 (Th1) and cytotoxic T cells in tumor immunesurveillance (Hamai et al., 2010; Matsui et al., 1999), it is tempting to test the hypothesis that PPARβ/δ may actually downregulate immuno surveillance through attenuating T helper type 1 (Th1) response. WIth the expanding knowledge of the relation between immune system and cancer and the role PPARβ/δ plays in modulating inflammation, it is almost certain that PPARβ/δ could regulate tumorigenesis through regulating the immune system. Conclusions In summary, PPARβ/δ could regulate tumorigenesis by targeting almost every hallmark of cancer and important hallmarks of cancer targeted by PPARβ/δ are summarized in table1. However, there are still a number of unsolved questions that need to be examined in order to fully understand the role of PPARβ/δ in regulating carcinogenesis. For example, what are the targets of action for some of the PPARβ/δ-independent functions of GW0742? What are the target genes that PPARβ/δ regulate to exert its anti-tumorigenic functions? Are there any other non-classical PPRE-independent regulations of transcription by PPARβ/δ that influences tumorigenesis? What is the status of endogenous ligands for PPARβ/δ 25

39 in different human cancers and is the difference in the level or types of endogenous ligands responsible for some of the opposite effects of PPARβ/δ on tumorigenesis? With the advent of new techniques and approaches such as next generation sequencing and metabolomics, hopefully these questions will be answered soon. Hallmarks of cancer Pro or no effects Anti Target genes Mechanism Target genes Mechanism Angiogenesis Apoptosis Represses p57 Kip2, Increases thrombospondin- 1 and CD36 Represses ILK and PDPK1 Increased morphologically immature and hyperplastic microvessels found surrounding tumors, causing a diminished blood flow and subsequent impaired tumor growth in the absence of PPARβ/δ. (Abdollahi et al., 2007; Muller-Brusselba ch et al., 2007) PPARβ/δ does not promote apoptosis and no increase of ILK and PDPK1 is found following ligand activation of PPARβ/δ and PPARβ/δ rather represses ILK and PDPK1 (Burdick et al., 2007; Marin et al., 2006; Yang et al., 2010) Represses VEGFR2 Increases ILK and PDPK1, represses PTEN PPARβ/δ agonists inhibited the formation of capillary-like structures and endothelial cell migration by downregulating endothelial VEGFR2 (Meissner et al., 2010) Ligand activation of PPARβ/δ leading to increased phosphorylation of AKT and subsequent increased anti-apoptotic and pro-survival capacity (Di-Poi et al., 2002) 26

40 Does not Increase VEGF Does not antagonize PPARγ activity Does not Increases expression Ligand activation of PPARβ/δ did not increase the expression of VEGF nor increase the phosphorylation of AKT in human cancer cell lines (Hollingshead et al., 2007) PPARβ/δ is unlikely to antagonize PPARγ activity since PPARβ/δ expression was similar between PPARγ-induced apoptosis resistant and sensitive cell lines (Foreman et al., 2009; He et al., 1999), NSAIDs does not decrease PPARβ/δ expression and neither NSAIDs or ligand activation of PPARβ/δ changed the level of ε (Foreman et al., 2011) Increases VEGF Antagonize PPARγ activity Increases expression Ligand activation of PPARβ/δ increased VEGF expression, causing increased phospho-akt level, which in turn promoted cell survival.(wang et al., 2006) PPARγ promoted apoptosis by decreasing survivin expression and increasing caspase-3 activity, and ligand activation of PPARβ/δ antagonized these effects.(wang et al., 2011) NSAIDs promotes apoptosis by repressing PPARβ/δ activity and expression (Liou et al., 2007; Wu and Liou, 2009) 27

41 Sustaining proliferative signaling Insensitivity to anti-growth signals Limitless replicative potential Increased glutathione peroxidase 1, thioredoxin 1, manganese superoxide PPARβ/δ activation reduces ROS and suppresses DNA damage induced Increases Ubc Increased secreted IL-1 receptor antagonist level Upregulates p21/p27 PPARβ/δ upregulated ubiquitin C and promoted ubiquitin-depend ent protein turnover of PKCα in mouse skin in response to TPA, resulting in a lower PKCα level, decreased Raf1, and MEK/ERK activities. (Kim et al., 2004a; Kim et al., 2005a). Dermal fibroblast PPARβ/δ inhibited the mitotic activity of keratinocytes via inhibition of the IL-1 signaling pathway by upregulation of IL-1 receptor antagonist, causing an autocrine decrease in IL-1 signaling pathways (Chong et al., 2009) BPS-mediated PPARβ/δ activation induces p21/p27 to inhibit aortic smooth muscle cell proliferation (Sue et al., 2009) 28

42 Chronic inflammation dismutase and heme oxygenase 1, Increased PTEN senescence through downregulating antioxidant genes or repressing PI3K-AKT pathway (Ham et al., 2012; Kim et al., 2011a; Kim et al., 2011b) Reduces IL1, IL6, TNF-α, MCP1 and CXCL7 Table 1. Key aspects of hallmarks of cancer that PPARβ/δ regulates Interfere with NF-κB signaling by sequestering p65 (See text for references) 1.4. PPARβ/δ and skin tumorigenesis In this section, I will discuss about the role PPARβ/δ plays in normal skin homeostasis and also about the anti-tumorigenic roles of PPARβ/δ in regulating chemically induced skin carcinogenesis PPARβ/δ regulates normal skin homeostasis Being the most abundant isoform of PPAR found in skin (Girroir et al., 2008a), It is not surprising that PPARβ/δ regulates normal skin homeostasis in at least two major ways: regulating skin wound healing process and inducing skin terminal differentiation. PPARβ/δ expression is upregulated in the keratinocytes at the wound edge of the damaged skin and its expression persisted until completion of the wound healing process (Michalik et al., 2001). Heterozygous PPARβ/δ mutant mice exhibited delayed completion of the healing process as compared to the wild-type animals (Michalik et al., 2001) and the delayed wound healing was due in part to a 29

43 severely reduced migration rate of keratinocytes (Michalik et al., 2001). In addition, PPARβ/δ upregulation occurred in the wound repair process after cutaneous injury rather than in the physiological renewal of the skin phase (Michalik et al., 2001). Furthermore, it was shown that PPARβ/δ upregulation is closely linked to necrosis and pro-inflammatory cytokines, such as interferon-γ and TNF-α, both of which can increase PPARβ/δ expression via an activator protein-1 (AP-1) binding site through the stress kinase pathway (Tan et al., 2001). TNF-α may also trigger the production of endogenous PPARβ/δ ligands, leading to an increase of PPARβ/δ transcriptional activity, which in turn confers an increased resistance of keratinocytes to TNF-α-induced apoptosis (Tan et al., 2001). This evidence suggests that PPARβ/δ is a key mediator of epidermal effects in wound healing by transducing the extracellular inflammatory signals into transcriptional regulation of gene expression leading to survival, migration and differentiation of keratinocytes (Kuenzli and Saurat, 2003). Perhaps the most uncontroversial functions of PPARβ/δ is the induction of terminal differentiation following ligand activation since it is supported by a plethora of evidence from multiple models, including keratinocytes, intestinal epithelium, osteoblasts, oligodendrocytes and monocytes, as well as various cancer models such as colon, breast and neuroblastoma cancers (Almad and McTigue, 2009; Aung et al., 2006; Boiteux et al., 2009; Borland et al., 2008; Burdick et al., 2007; Burdick et al., 2006; Di Loreto et al., 2007; Hollingshead et al., 2008; Kim et al., 2006a; Man et al., 2008; Marin et al., 2006; Matsuura et al., 1999; Nadra et al., 2006; Saluja et al., 2001; Schmuth et al., 2004; Still et al., 2008; Tan 30

44 et al., 2001; Varnat et al., 2006; Vosper et al., 2003; Werling et al., 2001; Westergaard et al., 2001; Yang et al., 2010). The induction of differentiation by ligand activation of PPARβ/δ depends on the increased expression of Angptl4 following ligand activation in keratinocytes (Pal et al., 2012). Importantly, ANGPTL4 stimulated the activation and binding of JUNB and c-jun to the promoter region of human involucrin and transglutaminase type 1 genes, respectively, thus facilitating the differentiation process (Pal et al., 2012) Mouse skin carcinogenesis models A major cause of skin cancer in humans is ultraviolet radiation (UV) radiation, which produces DNA lesions in affected skin. UV radiation produces DNA mutations in critical proto-oncogenes and tumor suppressor genes such as p53 which lead to skin cancer development and progression (Aszterbaum et al., 1999; Stratton et al., 2000). Additionally, industrial chemicals have been linked to the increased incidence of skin cancer in humans due to the ubiquitous mutagenic nature of some of these toxicants. Skin tumors can be divided into two broad types based on the ability of the tumor to metastasize (Stratton et al., 2000). Benign skin tumor is characterized by the inability of the tumor to invade surrounding tissue or metastasize, and includes papillomas. Malignant skin tumor is characterized by the ability of the tumor to invade surrounding tissue or metastasize, and includes basal cell carcinomas (BCC) and squamous cell carcinomas (SCC) (Stratton et al., 2000). Somewhat controversial is the definition of keratoacanthoma as its variants are clinically and histologically heterogenous 31

45 and while some consider the keratoacanthoma to be benign, others classify it as a subtype of squamous cell carcinoma (Ko, 2010; Mandrell and Santa Cruz, 2009). However, in mouse models, keratoacanthomas are though to be premalignant since they have a higher frequency of spontaneous malignant transformation to squamous cell carcinomas (Brown et al., 1998). To mimic the effects of UV and environmental carcinogens on skin carcinogenesis, two skin carcinogenesis mouse models have been developed. The UV-induced mouse skin carcinogenesis model usually utilizes SKH1 albino hairless mice, which have enhanced sensitivity to UV-induced tumorigenesis (Yarosh and Yee, 1990). Mice are exposed to repeated UV radiation and tumor growth is detected macroscopically. In contrast to the multiple stages of tumorigenesis by chemically-induced skin carcinogenesis, UV radiation is a complete carcinogen and therefore distinct stages of carcinogenesis cannot be distinguished with repeated exposures to UV radiation. UV radiation causes photochemical cellular damage, with UVB ( nm) being the most carcinogenic radiation. In addition to inducing DNA damage, UVB radiation can also disrupt tumor suppressor pro-apoptotic signaling pathways and suppress immune responses (Nghiem et al., 2001; Ullrich and Schmitt, 2000). Most human skin cancers that are caused by UV radiation from sunlight contain p53 mutations as the predominant mutation (Brash et al., 1991; Kanjilal et al., 1993; Tornaletti et al., 1993). In addition to mutations in the p53 gene, UV radiation also induces mutations in the ras proto-oncogene in certain rodent strains, especially mutations in Nras gene (Pierceall et al., 1992). 32

46 The chemically (two-stage)-induced skin carcinogenesis model is another widely used model that involves three stages including initiation, promotion and progression (Yuspa et al., 1976). Initiation is caused by an exposure to a sub-threshold dose of a carcinogen that causes mutation(s) in critical gene(s). Widely used initiators in the chemically-induced skin carcinogenesis model include 7, 12 dimethylbenz- anthracene (DMBA), benzopyrene, urethane (ethyl carbamate), and N-methyl-N -nitro-n-nitrosoguanidine (MNNG). MNNG can directly bind to DNA and cause alteration to bases, which results in mutation(s) following replication or transcription in the absence of DNA repair (Kartasheva and Bykorez, 1975; Ketkar et al., 1978). However, polyaromatic hydrocarbons (PAHs), such as DMBA, are biologically inert and require bio-activation into electrophilic metabolites. They are metabolized by cytochrome P450 monooxygenases and microsomal epoxide hydrolase that convert PAHs into highly reactive diol epoxides (Desai et al., 2002; DiGiovanni, 1989; Kondraganti et al., 2003), which then can covalently bind to exocyclic amino groups of guanine and adenine, forming stable adducts within DNA (Desai et al., 2002; DiGiovanni, 1989; Kondraganti et al., 2003). In the absence of DNA repair, the diol epoxide-dna adducts may become permanent mutations following DNA replication or transcription (Melikian et al., 1991). The Hras oncogene is the most critical and commonly mutated gene in chemically-induced skin carcinogenesis (Balmain et al., 1984; Quintanilla et al., 1986; Quintanilla et al., 1991) and over 90% of the papillomas induced by DMBA contained activating mutations in Hras gene with an A to T transversion at codon 61 (Balmain et al., 1984; Quintanilla et 33

47 al., 1986). Additionally, MNNG, an alkylating agent and carcinogen, also causes an activating mutation in the Hras gene in half of the papillomas in the skin carcinogenesis model (Quintanilla et al., 1986). The second stage of skin carcinogenesis is promotion, which is accomplished through repeated application of a non-mutagenic agent that stimulates proliferation. Tumor promotion results in clonal selection and expansion of initiated cells and subsequent tumor development (Hennings et al., 1993). The most commonly used tumor promoter in chemically-induced skin carcinogenesis is 12-O-tetradecanoylphorbol-13-acetate (TPA). TPA promotes tumor growth by activating the protein kinase C (PKC) pathway due to its functional analogy to diacylglyerol (DAG) (Slaga et al., 1982). In addition; TPA can promote tumor growth by its pro-inflammatory property (Slaga et al., 1982). A relatively small percentage of the papillomas that developed following tumor promotion progress to become malignant skin tumors and this is the progression stage. Tumor progression of benign papilloma to a malignant carcinoma is characterized by increased genetic instability in tumor cells and often involves additional mutations in other critical tumor suppressor and oncogenes (Yuspa and Poirier, 1988). Different mouse strains have been shown to display varying degree of sensitivity to the two-stage skin carcinogenesis bioassay. SENCAR mice have been shown to be the most sensitive to the chemically-induced skin carcinogenesis model (Hennings et al., 1981). C57BL/6 mice have a medium sensitivity among mice strains and show a resistance to malignant conversion of benign tumors (Slaga et al., 1982) To identify the nature and location of the target cell in Hras-induced skin 34

48 carcinogenesis, namely, the cell in skin that tumor is most likely to arise from; several skin specific Hras transgenic mice have been developed. While transgenic mice expressing Hras under the keratin 10 promoter where Hras is expressed mainly in superbasal cells of epidermis only developed benign papillomas at sites of promotional stimuli such as scratching, wounding or TPA treatment (Bailleul et al., 1990), transgenic mice expressing Hras under the keratin 5 promoter where expression was in the basal layer of epidermis developed spontaneous papilloma-like and keratoacanthoma-like tumors, and these keratoacanthoma-like tumors frequently underwent malignant conversion to squamous cell carcinomas (Brown et al., 1998). Since transgenic mice expressing Hras under keratin 5 promoter recapitulated the complete spectrum of events in the multistage skin carcinogenesis, this suggested that cells in basal layer rather than in the superbasal layer of epidermis is the tumor initiating cells (Brown et al., 1998) In addition to the in vivo skin carcinogenesis models, some in vitro skin cancer cell lines that contain Hras mutations at codon 61 (the same as the mutations in two stage bioassay) have also been developed. SP-1 and BP-4 cell lines were derived from pools of papillomas produced on SENCAR and BALB/c mouse skin, respectively, by initiation with DMBA and promotion with TPA (Strickland et al., 1988). Line 308 was derived from BALB/c mouse skin initiated in vivo with DMBA, culture of the epidermal cells, and selection of cells resistant to Ca 2+ -induced terminal differentiation (Strickland et al., 1988). Line LC14 was derived from untreated, cultured newborn BALB/c mouse primary epidermal cells, which 35

49 spontaneously developed resistance to Ca 2+ -induced terminal differentiation (Strickland et al., 1988). Furthermore, retroviral introduction of oncogenic Hras into normal mouse primary keratinocytes can transforme these normal keratinocytes into initiated keratinocytes by blocking their ability to undergo differentiation (Yuspa, 1985a; Yuspa, 1985b; Yuspa et al., 1985). Additionally, grafting a mixture of these initiated keratinocytes that express oncogenic Hras with normal dermal fibroblasts produces papillomas in recipient athymic nude mice (Greenhalgh et al., 1990; Harper et al., 1986; Roop et al., 1986; Yuspa et al., 1990). However, another study reported that co-grafting of Hras initiated keratinocytes with normal dermal fibroblasts inhibits malignant transformation in syngenic graft experiments (Dotto et al., 1988). Co-grafting of normal fibroblasts and established cancer cell lines such as 308, LC 14 and SP1 keratinocyte lines also produces papillomas and carcinomas in vivo (Strickland et al., 1988). Since Hras mutation is critical in the multistage skin carcinogenesis, these excellent ex vivo models provided flexible approaches for the study of Hras-induced skin carcinogenesis PPARβ/δ regulates skin carcinogenesis The first study suggesting that PPARβ/δ regulates epidermal cell proliferation was obtained by comparing epidermal cell proliferation in response to TPA treatment between wild-type and Pparβ/δ-null mice (Peters et al., 2000a). The hyperplastic response observed in the epidermis after TPA application was significantly greater in both the Pparβ/δ-null and Pparβ/δ +/ mice than in controls (Peters et al., 36

50 2000a, Michalik et al., 2001). Following studies utilizing chemically-induced carcinogenesis models showed that Pparβ/δ-null mice were more sensitive to DMBA-TPA induced skin carcinogenesis compared to wild-type mice (Kim et al., 2004a). Enhanced TPA-induced hyperplasia in Pparβ/δ-null mice was a result of reduced ubiquitin C, whose expression is regulated by PPARβ/δ (Kim et al., 2004a). PPARβ/δ promoted ubiquitin-dependent proteosome-mediated protein turnover of PKCα in mouse skin in response to TPA, resulting in a lower PKCα level, decreased Raf1, and MAPK/extracellular signal-regulated kinase activities and subsequent decreased cell proliferation (Kim et al., 2005a). However, selective ablation of PPARβ/δ in basal keratinocytes of adult mouse skin had no significant effects on DMBA/TPA-induced tumorigenesis (Indra et al., 2007), Therefore, PPARβ/δ might exert its tumor supressive activity in non-keratinocyte cells, and /or its expresssion in keratinocytes of the developing epidermis might prevent tumorigenesis (Indra et al., 2007). However, the targeting strategy for PPARβ/δ deletion in skin were unknown and western blot analysis showing the absence of PPARβ/δ following deletion was not available in this study, therefore, it is equally likely that the absence of effects on DMBA/TPA-induced tumorigenesis is due to inefficient deletion of PPARβ/δ in the skin. Following studies also showed that ligand activation of PPARβ/δ by a synthetic ligand GW0742 further attenuated chemically-induced carcinogenesis, which was due in part to induction of terminal differentiation and decreased secretion of inflammatory cytokines (Bility et al., 2008; Bility et al., 2010; Zhu et al., 2010). In addition, ligand activation also decreased cell proliferation of 308 and SP1 cells 37

51 through similar mechanisms (Bility et al., 2008; Bility et al., 2010). These findings clearly suggest that PPARβ/δ attenuates chemically-induced skin carcinogenesis. The hypothesis that PPARβ/δ regulates UV-induced skin carcinogenesis is still under investigating at this time Hypothesis PPARβ/δ is a nuclear receptor with relatively high expression observed in the skin (Girroir et al., 2008a). Previous studies showed that Pparβ/δ-null mice exhibited enhanced sensitivity to chemically induced skin carcinogenesis as compared to wild-type mice, including earlier onset of tumor growth, higher tumor multiplicity and larger average tumor size (Bility et al., 2008; Bility et al., 2010; Kim et al., 2004a; Kim et al., 2005a; Zhu et al., 1998). In addition, ligand activation of PPARβ/δ attenuated chemically induced skin carcinogenesis in wild-type mice, including delayed tumor onset, lower tumor multiplicity but having no effects on tumor size (Bility et al., 2008; Bility et al., 2010; Zhu et al., 2010). This was in part due to the induction of terminal differentiation and inhibition of DNA synthesis by ligand activation of PPARβ/δ (Bility et al., 2008; Bility et al., 2010; Zhu et al., 2010). Furthermore, ligand activation of PPARβ/δ inhibited cell proliferation and induced terminal differentiation in immortalized cancer cell lines including 308, SP1 and Pam212 (Bility et al., 2008). Additionally, TPA-induced inflammation in skin was lower in wild-type mice compared to similarly treated Pparβ/δ-null mice, suggesting a role of PPARβ/δ in the attenuation of TPA induced inflammation (Kim et al., 2004a; Peters et al., 2000a). Since over 90% of the tumors initiated by 38

52 DMBA contain mutations in the Hras proto-oncogene, it is very likely that PPARβ/δ inhibits epidermal cell proliferation and skin carcinogenesis via inhibition of oncogenic Ras signaling in a DMBA-TPA two stage carcinogenesis model. Furthermore, combining PPARβ/δ agonists with other chemotherapeutic agents could enhance the efficacy of attenuation of skin carcinogenesis by either agent alone. The focus of this study is to develop model systems to test this hypothesis, and to elucidate the mechanism by which PPARβ/δ regulates Hras signaling in the process of skin carcinogenesis. To elucidate the mechanism by which PPARβ/δ modulates Hras signaling in skin, primary keratinocytes with retrovirally introduced oncogenic Hras are used as the main model. Since a brief increase of cell proliferation caused by Hras will be followed by a cellular senescence response in primary keratinocytes (Vijayachandra et al., 2003), cell proliferation and senescence will be quantified in primary keratinocytes in the absence and presence of GW0742, a PPARβ/δ specific ligand. To search for the genes that PPARβ/δ regulates in response to an oncogenic Hras signaling, microarray analysis will be performed in both mock-infected and HRAS-expressing wild-type and Pparβ/δ-null keratinocytes with or without GW0742. In addition, potential differentially expressed genes between DMSO treated wild-type HRAS-expressing keratinocytes and GW0742 treated wild-type HRAS-expressing keratinocytes and between DMSO treated wild-type HRAS-expressing keratinocytes and DMSO treated Pparβ/δ-null HRAS-expressing keratinocytes will be verified using in vivo models. A two-stage skin carcinogenesis assay will be performed to validate differentially expressed 39

53 genes involved in cell proliferation and a one-stage skin carcinogenesis assay, where DMBA will be applied repeatedly without TPA promotion, will be used to examine different senescence phenotypes in vivo, since more senescent cells were observed using this approach as compared to two-stage bioassay (Sun et al., 2007). Due to the complexity of most cancers, cocktail strategies have been the most effect therapeutical approaches. Ligand activation of PPARβ/δ attenuates chemically induced skin carcinogenesis (Bility et al., 2008). In addition, COX-2 inhibition by NSAID also have proven to be an effective anti-tumorigenic therapy (Husain et al., 2002). A number of recent reports suggest that PPARβ/δ is an effector in COX2 signaling mediated promotion of colorectal cancer growth and invasion (Gupta et al., 2000; Wang et al., 2004). However, recent reports from our lab showed that PPARβ/δ and COX2 inhibitors attenuate skin and colon tumorigenesis via independent mechanisms (Hollingshead et al., 2008; Kim et al., 2006c). Since combining ligand activation of PPARβ/δ with COX2 inhibition by nimesulide had modest increased efficacy in the attenuation of chemically induced skin carcinogenesis in a chemotherapeutical model (Bility et al., 2010), it is hypothesized that combination of ligand activation of PPARβ/δ with COX2 inhibition might have an increased efficacy in the attenuation of chemically induced skin carcinogenesis in a chemopreventionl model. This hypothesis is tested using a two-stage carcinogenesis bioassay model and the results are illustrated in chapter four of my thesis. 40

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73 Chapter 2 PPARβ/δ crosstalks with E2F and attenuates mitosis in HRAS-expressing cells 2.1 Abstract The role of PPARβ/δ in Harvey sarcoma ras (Hras) expressing cells was examined. Ligand activation of PPARβ/δ caused a negative selection against cells expressing higher levels of the Hras oncogene by inducing a mitotic block. Mitosis-related genes that are predominantly regulated by E2F were induced to a higher level in HRAS-expressing Pparβ/δ-null keratinocytes as compared to HRAS-expressing wild-type keratinocytes. Ligand activated PPARβ/δ repressed expression of these genes by direct binding with p130/p107, facilitating nuclear translocation, and increasing promoter recruitment of p130/p107. These results demonstrate a novel mechanism of PPARβ/δ crosstalk with E2F signaling. Since co-treatment with a PPARβ/δ ligand and various mitosis inhibitors increases the efficacy of increasing G2/M arrest, targeting PPARβ/δ in conjunction with mitosis inhibitors could become a suitable option for development of new multi-target strategies for inhibiting RAS-dependent tumorigenesis. 2.2 Introduction Targeting peroxisome proliferator-activated receptors (PPARs) for the prevention and treatment of diseases is of great interest due to their ability to modulate many physiological functions (Akiyama et al., 2005; Grimaldi, 2005; Lee et al., 2003; Peters and Gonzalez, 2009; Peters et al., 2008). PPARβ/δ ligands can increase serum high density lipoprotein cholesterol concentration, improve insulin 60

74 resistance, increase fatty acid catabolism, and exert potent anti-inflammatory activities (Akiyama et al., 2005; Grimaldi, 2005; Kilgore and Billin, 2008; Lee et al., 2003; Lee et al., 2006; Oliver et al., 2001). There is also potential for targeting PPARβ/δ for the prevention and treatment of cancer. However, the role of PPARβ/δ in cancer remains controversial (reviewed in (Peters et al., 2011a; Peters and Gonzalez, 2009; Peters et al., 2008; Peters et al., 2012; Peters et al., 2011b)). The first evidence suggesting that PPARβ/δ modulates skin carcinogenesis was provided by the observation that Pparβ/δ-null mice exhibit enhanced epidermal hyperplasia in response to the tumor promoter 12-O-tetradecanoylphorbol-13-acetate (TPA) (Peters et al., 2000b). Consistent with this phenotype, exacerbated skin tumorigenesis is also found in Pparβ/δ-null mice following a two-stage chemical carcinogenesis bioassay (Kim et al., 2004b). Subsequent studies demonstrated that ligand activation of PPARβ/δ inhibits chemically-induced tumorigenesis (Bility et al., 2008; Bility et al., 2010; Zhu et al., 2011). One mechanism that may underlie these PPARβ/δ-dependent chemopreventive effects is modulation of epidermal cell proliferation through inhibition of PKCα/MAPK (Kim et al., 2004; Kim et al., 2005). Additionally, ligand activation of PPARβ/δ also induces terminal differentiation (Bility et al., 2008; Kim et al., 2006; Westergaard et al., 2001), which influence cell proliferation and skin tumorigenesis. Neoplastic conversion of normal cells to benign tumors and progression of benign tumors to adenomas and carcinomas is associated with over-expression, amplification, and homozygosity of oncogenic Hras (Malumbres and Barbacid, 61

75 2003). Chemicals can cause mutations in Hras in mouse skin tumors (Bizub et al., 1986), and examination of skin tumors produced by chemical carcinogens or ultraviolet light revealed that nearly all tumors contain an activated Hras oncogene (Balmain et al., 1984; Daya-Grosjean et al., 1993). Targeted introduction of oncogenic Hras into the epidermis of experimental animals can replace the initiation step resulting from exposure to a mutagenic chemical such as 7,12-dimethylbenz[a]anthracene (DMBA) in a two-stage chemical carcinogenesis model (Brown et al., 1986) and introduction of an Hras oncogene into normal mouse keratinocytes can also produce benign papillomas when grafted onto nude mice (Greenhalgh et al., 1990). Thus, there is strong evidence suggesting that activating the Hras oncogene through mutagenesis contributes to the mechanisms leading to neoplastic transformation during skin carcinogenesis. Previous work demonstrated that activation of PPARβ/δ attenuates skin tumorigenesis in a 2-stage chemical carcinogenesis bioassay and inhibits proliferation of cells with a mutation in the Hras gene (Bility et al., 2008). This suggests that PPARβ/δ could inhibit tumorigenesis through inhibition of oncogenic Hras signaling, which was examined in these studies. 2.3 Materials and methods Virus production The Hras retrovirus was generated from ψ2 producer cells as described previously (Roop et al., 1986). The virus titer was determined to be between 1 2 x 62

76 10 7 transforming units/ml using an NIH-3T3 focus-forming assay Plasmids pcmv-p130 and pcmv-e2f4 vectors were purchased from Origene (Rockville, MD). The pcmv-p107 vector was a kind gift from Dr. Liang Zhu (Zhu et al., 1993). The FLAG-p107, pgex-2t-p107 and pgex-2t-p130 vectors were kindly provided by Dr. Xavier Graña (Jayadeva et al., 2010). The psg5-pparβ/δ has been described previously (Girroir et al., 2008b). pcdna-flag-pparβ/δ was constructed by digesting ptnt-flag-pparβ/δ kindly provided by Dr. Gary Perdew from Penn State University with KpnI and NotI and ligated into KpnI and NotI linearized pcdna3.1 vector. The mouse Cdk1 promoter from -205 to + 57 was cloned using the following primers: forward primer: 5 -ATAGGTACCGGAAGGAAAACAGAGCTCAAGAG-3 ; reverse primer: 5 -ATACTCGAGCACACCGCAGTTCCGG-3. The PCR product was digested with KpnI and XhoI and ligated into KpnI and XhoI linearized pgl4.20 vector (Promega, Madison, WI). Site directed mutagenesis was performed to produce mutations in the following regions of the mouse Cdk1 promoter reporter construct (pgl4.20-mcdk1-promoter): distal E2F binding site: sense primer: 5 -GTTTCCGCTCCCTTTCGTAATCTGCGCTCCCAGGC-3 ; antisense primer: 5 -GCCTGGGAGCGCAGATTACGAAAGGGAGCGGAAAC-3 ; proximal E2F binding site: sense primer: 5 -GATCCCGGGAGCTTTAATATTGCGAGTTTGAAACTGC-3 ; antisense primer: 5 -GCAGTTTCAAACTCGCAATATTAAAGCTCCCGGGATC-3 ; CHR binding site: 63

77 sense primer: 5 -CTTTACCGCGGCGAGTCGACAACTGCTGGCACTCGG-3 ; antisense primer: 5 -CCGAGTGCCAGCAGTTGTCGACTCGCCGCGGTAAAG-3. Mutations were confirmed by direct sequencing and the following mutant reporter constructs were obtained: the mouse Cdk1 promoter reporter construct with a mutant distal E2F binding site (pgl4.20-mcdk1-distal E2F mutant), the mouse Cdk1 promoter reporter construct with a mutant proximal E2F binding site (pgl4.20-mcdk1-proximal E2F mutant) and the mouse Cdk1 promoter reporter construct with a mutant CHR binding site (pgl4.20-mcdk1-chr mutant) Cell culture Primary keratinocytes from newborn wild-type and Pparβ/δ-null mice were prepared and cultured as previously described (Dlugosz et al., 1995). Keratinocytes were infected with the Hras retrovirus for two days at an estimated MOI of Cells were subsequently cultured in control medium or medium containing GW0742 for up to nine days post-infection. 308 keratinocytes that contain an activated mutation in Hras (Strickland et al., 1988; Yuspa and Morgan, 1981) were cultured in low calcium medium as described previously (Bility et al., 2008; Bility et al., 2010) Cell proliferation assay Wild-type and Pparβ/δ-null keratinocytes were infected with an Hras encoding retrovirus at an estimated M.O.I of 3 for two days. Keratinocytes were then treated 64

78 with or without 1 µm GW0742 for another four days. Cell number was quantified using a Z1 Beckman Coulter. Alternatively, cells were seeded in 96 well plates and after seventy-two hours of treatment with or without 1 µm GW0742, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) in PBS was added to each well at the final concentration of 0.5 mg/ml and cells were incubated at 37 C for 1 hour. The media was removed to quantify optical density with a spectrophotometer at 560 nm Immunofluorescence analysis HRAS-expressing wild-type and Pparβ/δ-null keratinocytes treated with and without 1 µm GW0742 were cultured in chamber slides to ensure complete attachment. Cells were fixed in 2% formaldehyde in PBS for 15 minutes at room temperature followed by permeabilization with 100% methanol for 10 minutes at -20 C. Cells were then washed with PBS and incubated overnight with an anti-phospho-histone 3 (S10) antibody (Cell Signaling, Beverly, MA) at 4 C followed by incubation with an Alexa-488 conjugated secondary antibody (Cell Signaling, Beverly, MA) and a Cy3-conjugated anti-tubulin antibody (Sigma-Adrich, St. Louis, MO) for 1 hour at room temperature in the dark. Cells were then washed with PBS before incubation in 1 μg/ml Hoechst for 10 minutes at room temperature. Paraffin embedded sections from skin tumors were prepared from samples collected from a previously published study, from wild-type and Pparβ/δ-null mice, with and without topical application of GW0742 (Zhu et al., 2011). Sections were deparaffinized with xylene, rehydrated with 65

79 decreasing concentrations of ethanol followed by antigen retrieval by boiling in 10 mm sodium citrate buffer (ph 6.0). phospho-histone 3 S10 (ph3s10), TUBULIN or total DNA (Hoechst) was detected as described above. The mitotic index was calculated as the percentage of cells stained positive for ph3s10. For every sample a minimum of 1000 total cells were examined. Cells were immunostained with antibodies against ph3s10 and β-tubulin, and co-stained with Hoechst to visualize DNA, to identify mitotic cells at different phases and the distribution of cells in various phases of mitosis was determined using a previously described method (Moffat et al., 2006). For every sample a minimum of 4000 total cells were examined. For co-localization analysis, cells were prepared as above. The anti-p107/p130 antibody (Santa Cruz Biotechnology, Santa Cruz, CA), anti-pparβ/δ 8095 antibody (Girroir et al., 2008b) and anti-e2f4 antibody (Santa Cruz Biotechnology, Santa Cruz, CA) were conjugated with Alexa-647, Alexa-488 or Alexa-568 dye, respectively using the manufacturer s recommended protocol (Invitrogen, Carlsbad, CA). Representative photomicrographs were obtained with an Olympus Fluoview 1000 confocal microscope, using a 60X oil objective (NA: 1.35) at room temperature. Photomicrographs were acquired with FV10-ASW2.0 VIEWER software (Olympus) and deconvolved with Autoquant software (Media Cybernetics, Bethesda, MD). 66

80 2.3.6 Flow cytometry analysis Cells were stained with bromodeoxyuridine (BrdU) and/or propidium iodide (PI) and analyzed for cell cycle progression as previously described (Borland et al., 2011; He et al., 2008). Briefly, for BrdU staining, subconfluent cells were pulsed with 10 μg/ml bromodeoxyuridine (BrdU) for 1 h, trypsinized, washed with PBS and fixed in 70% ethanol overnight. Next day, cells were washed once with PBS and denatured by resuspending in 2M HCl followed by neutralization in 0.1 M Na 2 B 4 O 7 and stained with either a FITC-labeled anti-brdu monoclonal antibody (Phoenix Flow Systems, San Diego, CA) and propidium iodide (PI). For analysis of DNA content by PI alone, cells were trypsinized and washed with phosphate buffered saline (PBS) once before overnight fixation in 70% ethanol. Cells were then washed with PBS and stained with PI. Approximately 10,000 cells/sample were analyzed by flow cytometry to detect PI using a FC500 flow cytometer (Beckman Coulter, Miami Lakes, FL). The percentage of cells at each phase of the cell cycle ± S.D. was determined with FCS Express software. For anti-hras staining analysis, cells were trypsinized and washed with PBS once before fixation in 2% formaldehyde in PBS for 15 minutes at room temperature followed by permeabilization with 100% methanol for 10 minutes at -20 C. Cells were washed with PBS and incubated overnight with an anti-hras antibody (C20, Santa Cruz Biotechnology, Santa Cruz, CA) followed by incubation with an Alexa-488 conjugated secondary antibody (Cell Signaling, Beverly, MA) for one hour at room temperature in the dark. Approximately 5,000 cells/sample were analyzed by flow cytometry using a Coulter XL-MCL (Beckman Coulter, Miami 67

81 Lakes, FL). Pearson s second skewness coefficient, defined as 3 (mean relative HRAS intensity median relative HRAS intensity)/ standard deviation of relative HRAS intensity was calculated to determine the relative distribution of cells with varying levels of HRAS DNA microarray analysis Wild-type and Pparβ/δ-null keratinocytes were infected with Hras encoding retrovirus at an estimated M.O.I of 3 for four days before treatment with or without 1 μm GW0742 for 24 hours. For non-hras infected keratinocytes, cells were mock infected for four days before treatment with or without 1 μm GW0742 for 24 hours. Total RNA was isolated using TRIZOL reagent (Invitrogen, Carlsbad, CA) and purified with RNeasy Mini Kit (Qiagen, Valencia, CA). One hundred ng of total RNA per sample was prepared for analysis with GeneChip Mouse Gene 1.0 ST Array (Affymetrix, Santa Clara, CA) according to the manufacturer s instructions. The Robust Multichip Average (RMA) approach was used for normalization of microarray data using the R/Bioconductor package as previously described (Do and Choi, 2006). To identify genes that were significantly induced by HRAS, a false discovery rate (FDR) cut-off of 0.1 was used. The DAVID algorithm was used to functionally categorize genes involved in different biological process as previously described (Huang et al., 2009). Principle component analysis (PCA) was performed using the R/Bioconductor package. Data have been deposited in NCBI's Gene Expression Omnibus (GEO) database ( and are accessible through accession number 68

82 GSE Gene Set Enrichment Analysis (GSEA) The log 2 transformed normalized values of the microarray data were used for GSEA (Mootha et al., 2003; Subramanian et al., 2005). The phenotype data set was constructed by averaging the log 2 values of 62 mitosis related genes. The Pearson metric was chosen to rank the genes and the phenotype-permutation option was set to compute the enrichment scores. Sets of E2F target genes were from two published data sets in which E2F target genes were confirmed by chromatin immunoprecipitation (Ren et al., 2002; Xu et al., 2007). The EGR1 gene set was obtained from two confirmed EGR1 target genes databases (Fu et al., 2003; Svaren et al., 2000). The Sp1 gene set was obtained from the online Molecular Signature database ( Genes containing a GGGGCGGGGT motif within a -2 kb to 2 kb of the transcription start site were considered as SP1 target genes. For each analysis, the normalized enrichment score (NES) is indicated together with the corresponding FDR. Transcription factor binding sites were identified using in silico analysis with MatInspector software (Genomatix, Ann Arbor, MI). 69

83 2.3.9 Quantitative western blot analysis Cell lysates and supernatants used for western blots and immunoprecipitations were prepared as previously described (Borland et al., 2011; Girroir et al., 2008). Briefly, for total cell lysates, cells were lysed with either 25 mm MENG buffer supplemented with 150 mm NaCl, 1%Triton-X100, 30 mm sodium fluoride, 40 mm glycerophosphate, 10 mm sodium pyrophosphate, 2 mm sodium orthovanadate and a protease inhibitor cocktail or with RIPA buffer supplemented with 30 mm sodium fluoride, 40 mm glycerophosphate, 10 mm sodium pyrophosphate, 2 mm sodium orthovanadate and a protease inhibitor cocktail. The supernatant after centrifugation was used for immunoprecipitations or western blot analysis. The NE-PER Nuclear Protein Extraction Kit (Thermo Scientific, Rockford, IL) was used to isolate nuclear and cytosol protein. Equal amounts of cytosol and nuclear protein were used for western blot analysis. Western blot analysis using radioactive detection methods was performed as previously described (Girroir et al., 2008b). The primary antibodies used were: anti-phospho-retinoblastoma (RB; S780), anti-phospho-retinoblastoma (RB; S795), anti-phospho-retinoblastoma (RB; S807/811), anti-aurkb, anti-cenp-a and anti-chek1 (Cell Signaling, Beverly, MA), anti-hras, anti-cdk1, anti-cdk2, anti-cdk4, anti-cyclin B1, anti-cyclin D1, anti-cks1/2, anti-nek2, anti-h2afz, anti-birc5, anti-retinoblastoma (RB), anti-e2f1, anti-e2f4, anti-p107 and anti-p130 (Santa Cruz Biotechnology, Santa Cruz, CA), anti-lactic dehydrogenase (LDH; Jackson Immunoresearch, West Grove, PA) and anti-β-actin (Rockland, Gilbertsville, PA). The anti-pparβ/δ antibody has been previously described (Girroir et al., 2008b). 70

84 RNA isolation and quantitative real-time PCR (qpcr) analysis Total RNA was isolated from cell lines or tumor samples using TRIZOL reagent (Invitrogen, Carlsbad, CA). Reverse transcription and qpcr was performed as previously described (Palkar et al., 2010). The relative level of mrna was normalized to that of glyceraldehyde 3-phosphate dehydrogenase (Gapdh) or 18s RNA levels Chromatin immunoprecipitation (ChIP) ChIP was performed as previously described (Palkar et al., 2010). Briefly, cells were cross-linked with formaldehyde (1%) for 10 minutes at room temperature and then glycine was added to stop the cross-linking reaction. Cells were washed with ice-cold PBS, incubated in lysis buffer for 30 minutes, and cell debris removed by centrifugation. The chromatin was sonicated to generate DNA fragments with a range of bp. Chromatin was pre-cleared with blocked protein A/G slurry (Santa Cruz Biotechnology, Santa Cruz, CA), and then equal amounts of pre-cleared chromatin was immunoprecipitated with one of the following antibodies: anti-p107, anti-p130, anti-e2f4, anti-e2f1 (Santa Cruz Biotechnology, Santa Cruz, CA) or acetylated histone 4 (Millipore, Temecula, CA). Rabbit IgG was used as a negative control. qpcr was performed to determine relative enrichment of specific proteins in different promoter regions and ubiquitin C genomic DNA was used for normalization. The following primers were used for ChIP analysis: Cdk1 proximal E2F binding site forward primer: 5 -AGCTCAGCTCTGATTGGCTCCTTT-3 ; reverse primer: 71

85 5 -TTTCAAACTCGCCGCGGTAAAGC-3 ; Cdk1 distal E2F binding site forward primer: 5 -AAACAGAGCTCAAGAGTCAGTTGGCG-3 ; reverse primer: 5 -GCAGAGCGCGAAAGGGAGCGGAAA-3 ; E2f1 E2F binding site forward primer: 5 -GGCCAATGGAGGAGGCGTT-3 ; reverse primer: 5 -TGCAAAGTCCGGGCCACTT-3 ; Chek1 E2F binding site forward primer: 5 -TTTACGGCAGAGGTGTGCGCTTT-3 ; reverse primer: 5 -TTCTCACCAAGCAGTCCTTTGCCA-3 ; Ubc sequence forward primer: 5 -CCAGTGTTACCACCAAGAAGGTCA-3 ; reverse primer: 5 -CCATCACACCCAAGAACAAGCACA ChIP-re-ChIP assay For improved cross-linking of proteins that are not directly bound to DNA, a modified cross-linking approach was used (Fujita and Wade, 2004). Briefly, cells were incubated with 5 mm dimethyl dithiobispropionimidate (DTBP) on ice for 30 minutes and the reaction was stopped by adding 100 mm Tris-HCl, ph 8.0 and 150 mm NaCl. A second cross-linking was then performed with 1% formaldehyde for 10 minutes at room temperature and stopped by the addition of glycine to the final concentration of M. The remaining steps were essentially the same as the ChIP protocol described above with the following exceptions. For the first ChIP, sonicated chromatin was incubated with 2 µg of the following antibodies: control rabbit IgG, anti-pparβ/δ (8095) antibody (Girroir et al., 2008b) or anti-e2f4 that was conjugated to agarose beads (Santa Cruz Biotechnology, Santa Cruz, CA). After elution with extraction buffer (0.1 M NaHCO 3, 1% SDS), samples were 72

86 diluted with buffer (20 mm Tris-HCl, 150 mm NaCl, 2 mm EDTA, 1% Triton-X100) and subject to a second immunoprecipitation with the same antibody (rabbit IgG, anti-pparβ/δ or anti-e2f4) for single pull-down or the anti-pparβ/δ antibody after the first E2F4 antibody pull down Immunoprecipitation assay Standard immunoprecipitations were performed as previously described (Girroir et al., 2008b). Briefly, Cells were lysed in 25 mm MENG buffer supplemented with 150 mm NaCl, 1%Triton-X100, 30 mm sodium fluoride, 40 mm glycerophosphate, 10 mm sodium pyrophosphate, 2 mm sodium orthovanadate and a protease inhibitor cocktail. The supernatant were precleared with pre-blocked protein A/G agarose beads before incubation with primary antibody captured protein A/G agarose beads overnight at 4 C. The resin was washed three times with lysis buffer and 3X SDS loading buffer was added to the washed resin and the supernatant was resolved using SDS-PAGE. For immunoprecipitations involving FLAG-tagged proteins, FLAG matrix gel (Sigma-Adrich, St. Louis, MO) was used to pull down FLAG-tagged proteins. After three washes, co-immunoprecipitates were eluted with releasing buffer (10 mm Tris-HCl, ph 7.5, 150 mm NaCl, 1% Triton-X100) containing 100 µg/ml FLAG peptide (Sigma-Adrich, St. Louis, MO). The eluted precipitates were then resolved with SDS-PAGE. For sequential immunoprecipitations, cell lysates were first subject to anti-flag pull-down and primary precipitates were eluted as described above. Eluted co-immunoprecipitates were diluted with lysis buffer and subject to second immunoprecipitation. For immunoprecipitations involving in vitro translated 73

87 proteins, E2F4, PPARβ/δ and p130 were in vitro translated with TNT Quick Coupled Transcription/Translation kit (Promega, Madison, WI). In vitro translated proteins were mixed with each other in the absence or presence of 1 µm GW0742 at 30 C for 30 minutes. The proteins were then diluted with 25 mm MENG buffer supplemented with 150 mm NaCl, 1%Triton-X100 and phosphatase and protease inhibitors before immunoprecipitation as described above Luciferase reporter assay 308 cells or primary keratinocytes that were either mock-infected or HRAS-expressing were transiently transfected with equal amount of endotoxin free pgl4.20, pgl4.20-mcdk1-promoter, pgl4.20-mcdk1-distal-e2f mutant, pgl4.20-mcdk1-proximal-e2f mutant or pgl4.20-mcdk1-chr mutant constructs and pcmv-renilla as described above. Forty-eight hours after transfection, cells were treated with DMSO or 1 µm GW0742 for 24 hours. Cells were lysed with 1X passive lysis buffer (Promega, Madison, WI) and luciferase activity measured with a luminometer In vitro kinase assay Two hundred ng of GST-p130 or GST-p107 protein was incubated with 500 ng of recombinant PPARβ/δ (ProteinOne, Rockville, MD) or dilution buffer in the presence of 50 mm Tris-HCl, ph 7.5, 150 mm NaCl and 1 mm EDTA with or without 1 µm GW0742 on ice for 30 minutes. After this step, 200 ng of a 74

88 CDK4/CYCLIN D1 complex (Invitrogen, Carlsbad, CA) and a mixture of 100 μm cold and 32 P-γ-ATP was added in the presence of 1X kinase assay buffer (50mM Tris-HCl, ph 7.5, 10 mm MgCl 2, 1 mm EDTA, 2 mm DTT, 40 mm β-glycerophosphate, 20 mm ρ-nitrophenylphosphate, 0.1 mm sodium vanadate and 0.01% Brij 35). The reaction was performed at 30 C for 15 minutes and the reaction was stopped by adding 15 μl of a 3X SDS loading buffer. The presence of phosphorylated p130 or p107 was detected by autoradiography and the presence of total p130, p107 was detected by western blot using an anti-gst antibody (Santa Cruz Biotechnologies, Santa Cruz, CA). For this assay, enhanced chemilluminesence (ECL) was used to detect proteins. The presence of recombinant PPARβ/δ was detected by western blot using an anti-pparβ/δ antibody (Abcam, Cambridge, MA). 2.4 Results Ligand activation of PPARβ/δ induces G2/M arrest causing selection against high HRAS-expressing cells The effect of ligand activation of PPARβ/δ was examined using primary mouse keratinocytes expressing activated HRAS (Roop et al., 1986). Cell proliferation was greater in HRAS-expressing Pparβ/δ-null cells as compared to HRAS-expressing wild-type cells and ligand activation of PPARβ/δ inhibited proliferation of HRAS-expressing wild-type cells (Figure 2-1A). This effect was due to a PPARβ/δ-dependent G2/M phase block (Figure 2-1B-E). This change in 75

89 Figure 2-1. Ligand activation of PPARβ/δ attenuates cell proliferation by inducing G2/M arrest. (A-C) HRAS-expressing wild-type and Pparβ/δ-null keratinocytes were treated with or without 1 µm GW0742 for 4 d. Cell number was quantified daily. Cell cycle analysis was performed after 3 d of treatment. (D, E) HRAS-expressing wild-type and Pparβ/δ-null keratinocytes were treated with or without 1 µm GW0742 for 9 d and cell cycle analysis was performed. For all datasets, N = 3-4 independent samples per treatment group. Values represent the mean ± SEM. *significantly different than wild-type vehicle control (DMSO), P #significantly less than GW0742-treated wild-type, P

90 Figure 2-2. Ligand activation of PPARβ/δ induces G2/M arrest causing selection against high HRAS-expressing cells. (A) Western blot analysis of HRAS-expressing keratinocytes 5 d or 11 d post-infection. Expression levels were normalized to β-actin and are presented as fold change relative to control DMSO. (B) HRAS-expressing wild-type and Pparβ/δ-null keratinocytes were treated with or without 1 µm GW0742 for 3 d and qpcr was performed to quantify copy number of genomic Hras DNA. (C) Flow cytometric analysis using 77

91 anti-hras antibody was performed and Pearson s second skewness coefficient for HRAS intensity was calculated. (D) Mock-infected or HRAS-expressing keratinocytes with increasing M.O.I. were treated with or without 1 µm GW0742 for 24 or 72 h and Hras mrna expression was measured by qpcr. (E) Cells were infected with an Hras retrovirus with increasing M.O.I. and after 3 d of culture with or without 1 µm GW0742, an MTT assay was performed. (F) HRAS-expressing keratinocytes with increasing M.O.I. were treated with or without 1 µm GW0742 for 3 d and the percentage of cells in different phases of the cell cycle was determined by flow cytometry after PI staining. (G) HRAS-expressing keratinocytes were treated with or without 1 µm GW0742, 5 nm paclitaxel or 10 nm paclitaxel for 3 d. Flow cytometric analysis of HRAS intensity in HRAS-expressing keratinocytes were performed as above for (C). For all datasets, N = 3-4 independent samples per treatment group. Values represent the mean ± SEM. *significantly different than wild-type vehicle control (DMSO), P Values with different superscripts are different, P proliferation was not due to altered apoptosis (data not shown). Surprisingly, HRAS expression was lower in HRAS-expressing wild-type cells following treatment with GW0742 as compared to controls but this effect was not found in HRAS-expressing Pparβ/δ-null cells, whose expression of HRAS was higher as compared to wild-type cells (Figure 2-2A). Since it is unlikely that PPARβ/δ regulates the viral promoter driving HRAS expression, the hypothesis that the reduced expression of HRAS was due to selection against cells expressing higher levels of HRAS was examined. Indeed, ligand activation of PPARβ/δ decreased the relative copy number of integrated viral Hras DNA and the percentage of cells with high expression of HRAS in HRAS-expressing wild-type cells; effects not found in HRAS-expressing Pparβ/δ-null cells (Figure 2-2B,C). This was consistent with reduced expression of Hras mrna observed in HRAS-expressing wild-type cells (Figure 2-2D). Additionally, PPARβ/δ-dependent inhibition of Hras mrna expression occurred sooner, and the magnitude of this effect was greater, with increasing level of HRAS (Figure 2-2D). The efficacy for inhibiting cell proliferation 78

92 by ligand activation of PPARβ/δ was greater with increased HRAS expression (Figure 2-2E). This also shows that there is a range of HRAS required to increase cell proliferation and above this range leads to inhibition of proliferation in wild-type keratinocytes; an effect not found in Pparβ/δ-null keratinocytes (Figure 2-2E). In addition, as the level of HRAS increased, the magnitude of PPARβ/δ-dependent increase in G2/M arrest was greater (Figure 2-2F). Collectively, these data suggests that ligand activation of PPARβ/δ selects against cells with higher expression of activated HRAS. Whether the G2/M arrest directly causes selection against cells with higher expression of HRAS was examined by quantifying relative HRAS expression after treatment with a known mitosis inhibitor. Similar to what is observed with ligand activation of PPARβ/δ, inhibition of the G2/M phase with paclitaxel caused selection against cells with higher expression levels of HRAS (Figure 2-2G). This suggests that G2/M arrest resulting from ligand activation of PPARβ/δ causes selection against cells expressing higher levels of HRAS Inhibition of mitosis by ligand activation of PPARβ/δ in HRAS-expressing cells A G2/M phase arrest could be mediated by a block in mitosis, which was examined in the following experiments by quantifying the mitotic index and comparing the effect of mitosis inhibitors. Ligand activation of PPARβ/δ markedly reduced the mitotic index in HRAS-expressing wild-type cells with a higher level of HRAS, which was reflected by a PPARβ/δ-dependent increase in the cells at the 79

93 Figure 2-3. Ligand activation of PPARβ/δ inhibits mitosis of HRAS-expressing keratinocytes. Wild-type and Pparβ/δ-null keratinocytes were infected with an Hras retrovirus at an estimated M.O.I of 3 or 12 for 2 d and then treated with or without 1 µm GW0742 for another 3 d. (A-left panel) Mitotic index. (A-right panel) The distribution of cells in different mitotic phases. (B-D) HRAS-expressing wild-type and Pparβ/δ-null keratinocytes were cultured for 4 d and then treated with paclitaxel for 24 h. (B) Cells were immunostained to determine the mitotic index. (C) Representative photomicrographs of immunostained keratinocytes. Arrowheads indicate cells with a metaphase plate on a bipolar spindle. A significant number of HRAS-expressing Pparβ/δ-null keratinocytes showed fully aligned chromosomes at metaphase plate after 20 nm paclitaxel treatment. Scale bar = 10 µm. (D) The distribution of cells in different mitotic phases was determined as described in Materials and methods. For all datasets, N = 3 independent samples per treatment group. Values represent the mean ± SEM. *significantly different than wild-type vehicle control (DMSO), P

94 G2/M boundary and a decrease in cells at metaphase, anaphase and telophase (Figure 2-3A). A lower percentage of cells at the G2/M boundary and a higher percentage of cells in metaphase, anaphase and telophase were observed in Pparβ/δ-null counterparts (Figure 2-3A). Blocking mitosis at prometaphase with paclitaxel caused a greater increase in the mitotic index in Pparβ/δ-null cells as compared to wild-type cells (Figure 2-3B). While paclitaxel effectively blocked HRAS-expressing wild-type cells in prometaphase, a number of HRAS-expressing Pparβ/δ-null cells proceeded to metaphase following treatment with paclitaxel (Figure 2-3C,D). Enhanced sensitivity to paclitaxel (a microtubule stabilizer that blocks mitosis)-induced inhibition of cell proliferation was found in HRAS-expressing Pparβ/δ-null cells as compared to wild-type cells (Figure 2-4A). This is consistent with a higher level of mitosis (Figure 2-3A) and enhanced proliferation in HRAS-expressing Pparβ/δ-null keratinocytes (Figure 2-1A). To examine the effect of ligand activation of PPARβ/δ on mitosis entry, the cells were synchronized at the G2 phase with RO-3306 (a CDK1 inhibitor) and then released into nocodazole to block at prometaphase in the presence or absence of GW0742. Ligand activation of PPARβ/δ decreased the mitotic index only in HRAS-expressing wild-type cells (Figure 2-4B,C). Further, the mitotic index was greater after release from the G2/M boundary in HRAS-expressing Pparβ/δ-null cells as compared to wild-type cells (Figure 2-4B,C). While the majority of HRAS-expressing wild-type cells released from the G2 block and treated with GW0742 remained in the G2/M boundary, a high percentage of HRAS-expressing Pparβ/δ-null cells proceeded to prophase and prometaphase as compared to 81

95 Figure 2-4. Ligand activation of PPARβ/δ delays mitosis entry in HRAS-expressing keratinocytes. (A) HRAS-expressing keratinocytes were treated with the indicated concentration of paclitaxel for 24 h and cell proliferation determined by MTT assay. (B, C) HRAS-expressing keratinocytes were synchronized at the G2 phase by treatment with RO-3306 for 36 h. Cells were then either maintained in RO-3306 or released from the G2 phase block and cultured with nocodazole with or without 1 µm GW0742 for 12 or 24 h. Representative photomicrographs and the mitotic index, cells in G2/M boundary and different mitotic phases were determined. For all datasets, N = 3 independent samples per treatment group. Values represent the mean ± SEM. *significantly different than wild-type vehicle control (DMSO), P

96 Figure 2-5. Ligand activation of PPARβ/δ increased cells with polyploidy DNA. Representative DNA histograms from HRAS-expressing keratinocytes treated with and without (A) 1 µm GW0742 or (B) 10 nm paclitaxel. The percentage of cells with polyploidy DNA is shown in the far right panel. For all datasets, N = 3 independent samples per treatment group. Values represent the mean ± SEM. *significantly different than wild-type controls, P controls (Figure 2-4C). Since it is known that keratinocytes in the G2/M state can exhibit polyploidy (Zanet et al., 2010), this was examined in HRAS-expressing cells. Ligand activation of PPARβ/δ increased cells with polyploidy DNA concomitant with an increase in cells at the G2/M block only in HRAS-expressing wild-type cells (Figure 2-5A). Similarly, a markedly greater increase in cells with polyploid DNA was found in HRAS-expressing wild-type, but not Pparβ/δ-null cells by treatment with paclitaxel (Figure 2-5B). 83

97 2.4.3 Ligand activation of PPARβ/δ decreases expression of E2F target genes that regulate mitosis in HRAS-expressing keratinocytes Microarray analysis was performed to identify potential genes that could regulate mitosis through PPARβ/δ. Principle Component Analysis showed that the major difference in gene expression profiles was due to expression of HRAS (Figure 2-6A). Differences in gene expression were markedly larger between HRAS-expressing wild-type and Pparβ/δ-null cells compared to control cells and the effect of ligand activation was also PPARβ/δ-dependent (Figure 2-6A). Gene Ontology analysis showed significant enrichment of HRAS-induced genes that regulate chromosome condensation and mitotic cell cycle in both genotypes, and the enrichment score was much higher in HRAS-expressing Pparβ/δ-null cells as compared to wild-type cells (Figure 2-6B). Eighty-two mitosis-related genes induced by HRAS in either wild-type or Pparβ/δ-null cells were identified by this analysis. Expression of sixty-two of these genes was repressed by ligand activation of PPARβ/δ in HRAS-expressing wild-type cells but not Pparβ/δ-null cells (Figure 2-6C). Further, the fold-induction caused by HRAS was greater in Pparβ/δ-null cells as compared to wild-type cells (Figure 2-6C). Changes in expression of eighteen genes selected based on microarray and bioinformatic analysis was verified by qpcr including Cdk1, H2afz, Chek1 and Cenpa (Figure 2-7). Western blot analysis of HRAS-expressing cells also showed PPARβ/δ-dependent repression of CDK1, CYCLIN B1, H2AFZ, CHEK1, CENPA and NEK2 by ligand activation, and these effects were not due to changes in cell cycle distribution (Figure 2-8A). The PPARβ/δ-dependent repression of CYCLIN 84

98 Figure 2-6. Ligand activation of PPARβ/δ decreases expression of genes that modulate mitosis in HRAS-expressing keratinocytes. Microarray analysis was performed using control or HRAS-expressing wild-type or Pparβ/δ-null keratinocytes. (A) PCA of normalized microarray data. (B) Differential enrichment of genes involved in different biological processes was determined by genotology analysis by DAVID. The criterion for inclusion in analysis was a minimum of 1.3-fold change by HRAS. (C) A significant enrichment (82 genes) for mitosis and chromosome condensation regulators in HRAS-expressing Pparβ/δ-null keratinocytes was found by DAVID. Ligand activation of PPARβ/δ repressed induction of 62 of the 82 mitosis genes in HRAS-expressing wild-type but not Pparβ/δ-null keratinocytes. B1 persisted (Figure 2-8B). Bioinformatic analysis of the promoters of the sixty-two mitosis related genes showed that while PPREs were not found, several common regulatory elements were present in most genes, including E2F, SP1, and EGR (Figure 2-9A). Since E2F is known to regulate mitosis (Ishida et al., 2001; Muller et al., 2001), the 85

99 Figure 2-7. Quantitative real-time PCR confirming decreased expression of genes that modulate mitosis in HRAS-expressing keratinocytes by ligand activation of PPARβ/δ. Confirmation of changes in mrna expression of mitosis-related genes regulated by ligand activation of PPARβ/δ in HRAS-expressing keratinocytes. qpcr analysis was performed 24 hours after treating HRAS-expressing cells with 1 µm GW0742. For all datasets, N = 3 independent samples per treatment group. Values represent the mean ± SEM. *significantly different than wild-type HRAS controls, P

100 Figure 2-8. Quantitative western blot analysis confirming decreased expression of genes that modulate mitosis in HRAS-expressing keratinocytes by ligand activation of PPARβ/δ. (A) Western blot analysis of mitosis-related genes regulated by ligand activation of PPARβ/δ in HRAS-expressing keratinocytes. Protein expression is compared between control cells (No Hras) or HRAS-expressing wild-type (WT) or Pparβ/δ-null (KO) cells. Control HRAS-expressing cells were obtained 2 days post-infection (Day2). The effect of ligand activation of PPARβ/δ was examined by treating HRAS-expressing cells 2 days post-infection with either DMSO (D) or 1 µm GW0742 (G) for hours (Day3, Day4, respectively). Inset represents distribution of cell cycle phases obtained 24 hours post-ligand treatment. (B) Western blot analysis of CDK4, CYCLIN D1, and CYCLIN B1 from HRAS-expressing keratinocytes treated with or without 1 µm GW0742 for 9 d. For all blots, hybridization signals for each protein was normalized to that of ACTIN and are presented as fold change relative to control DMSO. For all datasets, N = 3 independent samples per treatment group. Values represent the mean ± SEM. *significantly different than wild-type HRAS controls, P Values with different superscripts are different, P

101 Figure 2-9. PPARβ/δ regulates E2F target genes in HRAS-expressing keratinocytes. (A) Promoter transcription factor binding sites analysis was performed to identify common transcription factors that regulate expression of the sixty-two differentially expressed mitosis-related genes modulated by PPARβ/δ. The P values indicate the relative significance of promoter enrichment for each respective transcription factor (B) The overlap of sixty-two mitosis-related genes and two ChIP-validated E2F target gene databases is presented in the Venn diagram. (C) Genes sets described in (B) were analyzed by Gene Set Enrichment Analysis (GSEA). The gene set of DNA replication and DNA repair was a subset of E2F target genes from Ren et al database. 88

102 Figure PPARβ/δ regulates E2F1 in HRAS-expressing keratinocytes. (A) qpcr validation of E2f1 and E2f4 expression (B) Western blot analysis of mock-infected (No Hras) or HRAS-expressing keratinocytes treated with or without GW0742 two to four days post-infection. Expression levels were normalized to β-actin and are presented as fold change relative to control DMSO. For all datasets, N = 3 independent samples per treatment group. Values represent the mean ± SEM. *significantly different than wild-type HRAS controls, P sixty-two genes were compared with two ChIP-confirmed E2F target gene databases (Ren et al., 2002; Xu et al., 2007). Twenty-two of these genes were also found to be E2F target genes based on this analysis (Figure 2-9B). Gene Set Enrichment Analysis (GSEA) revealed that E2F target genes were regulated similarly as the mitosis related genes, including genes involved in DNA repair and synthesis (Figure 2-9C); SP1 and EGR were ruled out as being central to this regulation (data not shown). E2F1 is an activator E2F that up-regulates expression of target genes whereas E2F4 is a repressor form of E2F that represses expression of target genes (Chen et al., 2009). PPARβ/δ-dependent 89

103 repression of E2F1 was observed following ligand activation (Figure 2-10A,B), consistent with the fact that E2F1 is auto-regulated (Johnson et al., 1994). Expression of E2F4 was not changed in response to ligand activation of PPARβ/δ (Figure 2-10A,B). The observed change in expression of mitosis-related genes was not mediated by altered phosphorylation of retinoblastoma (RB), because no change in phospho-rb was observed following ligand activation of PPARβ/δ (Figure 2-10B). This suggests that PPARβ/δ-dependent modulation of mitosis-related gene expression is downstream of RB. Consistent with the finding that ligand activation of PPARβ/δ in 308 cells causes G2/M arrest (Bility et al., 2010), repression of E2F target genes that regulate mitosis including CYCLIN B1, CHEK1, CDK1 and H2AFZ was also observed following ligand activation of PPARβ/δ in these cells (Figure 2-11A). This is important to note because 308 cells have an activated Hras mutation, in contrast to the keratinocyte model of viral HRAS transformation. Interestingly, the observed repression was greatest in cells with higher confluence when E2F activity was highest (Figure 2-11C). Additionally, expression of HRAS was repressed in response to ligand activation of PPARβ/δ but only after 72 hours of treatment when the cells were at higher confluence (Figure 2-11A). Ligand activation of PPARβ/δ in 308 cells for only 24 hours in confluent cells caused significant repression of E2F target genes that regulate mitosis with no apparent change in HRAS expression indicating that reduced HRAS expression does not mediate these changes (Figure 2-11A). As found in HRAS-expressing keratinocytes, expression of E2F1 was repressed by ligand activation of PPARβ/δ 90

104 Figure Ligand activation of PPARβ/δ inhibits expression of pro-mitotic proteins in 308 cells. 308 cells were cultured for up to 72 h and treated with GW0742 to activate PPARβ/δ for varying periods of time illustrated by the arrows. Western blot analysis of (A) mitosis related proteins or (B) RB, E2F1 and E2F4 in 308 cells treated as described above. Expression levels were normalized to β-actin and are presented as fold change relative to control DMSO. (C) 308 cells were cultured for 24, 48 or 72 after transfection with an E2F-luciferase reporter construct. Luciferase activity was measured and normalized to renilla as an internal control. in 308 cells (Figure 2-11B). Moreover, examination of phospho-rb showed that PPARβ/δ-dependent modulation of mitosis genes in 308 cells is downstream of RB (Figure 2-11B). 91

105 2.4.4 Ligand activation of PPARβ/δ represses CDK1 and E2F1 by increasing recruitment of p107/p130 to E2F4 binding sites The molecular mechanisms of repression of E2F target genes following ligand activation of PPARβ/δ was examined next. The decreased expression of E2F target genes could be caused by an increase of repressor E2F4 activity as the repressor E2F4 is known to form a complex with RB/p107/p130 to repress target gene expression (Chen et al., 2009). The nuclear to cytosol ratio of p130 (hypophosphorylated), p107, E2F4 and PPARβ/δ was increased by ligand activation of PPARβ/δ in 308 cells (Figure 2-14A) and in HRAS-expressing wild-type but not Pparβ/δ-null cells (Figure 2-12A,B). These effects were not observed in mock-infected keratinocytes (Figure 2-12B). Since the CDK1/CYCLIN B1 complex is known to be critical for mitosis entry, and ligand activation of PPARβ/δ repressed HRAS-induced CDK1 expression, the promoter occupancy of E2Fs was examined in the distal E2F1 (activator) and proximal E2F4 (repressor) binding sites of the CDK1 promoter (Zhu et al., 2004). Ligand activation of PPARβ/δ caused a reduction in acetylated histone 4 in both the E2F1 and E2F4 binding sites and decreased promoter occupancy of E2F1 on the E2F1 binding site in HRAS-expressing wild-type but not Pparβ/δ-null cells (Figure 2-13A). While no occupancy of p130 or p107 to the E2F1 binding site was found (data not shown), promoter occupancy of p130 to the E2F4 binding site was increased following ligand activation of PPARβ/δ in HRAS-expressing wild-type but not Pparβ/δ-null keratinocytes (Figure 2-13A). E2F4 promoter occupancy was evident on the E2F4 binding site; ligand activation of PPARβ/δ did not alter this occupancy 92

106 Figure Ligand activation of PPARβ/δ increases nuclear p107/130 protein in HRAS-expressing cells. Wild-type and Pparβ/δ-null cells were mock infected (B) or Hras-infected (A, B) for 2 d. Cells were cultured with or without 1 µm GW0742 for 24 h. Western blot analysis of cytosol (C) and nuclear (N) extracts from HRAS-expressing keratinocytes. Expression levels were normalized to β-actin. The average ratio of nuclear to cytoplasmic protein (N/C) is shown. Values represent the mean ± SEM. *significantly different than wild-type HRAS DMSO controls, P (Figure 2-13A). None of these changes were observed in mock-infected keratinocytes (Figure 2-13A). Similar effects were also noted in 308 cells (Figure 2-14B). Combined, these observations are consistent with the notion that ligand activation of PPARβ/δ represses HRAS-induced expression of CDK1 by repressing E2F1 activator activity and also increasing E2F4/p130 repressor activity. While the change in occupancy of E2F4/p130 on the CDK1 promoter following ligand activation of PPARβ/δ are associated with only a 37% decrease in expression of CDK1 protein, the stoichiometry of transcriptional and translational proteins required to mediate this repression is unknown and could involve multiple biological factors in addition to E2F4/p130 binding. Ligand activation of PPARβ/δ also caused a reduction in acetylated histone 4 and increased promoter occupancy of p130 and p107 on the E2F binding site of 93

107 FIgure Ligand activation of PPARβ/δ represses CDK1 and E2F1 by increasing recruitment of p107/130 and represses CHEK1 by decreasing recruitment of E2F1 to respective promoters in HRAS-expressing keratinocytes. Promoter occupancy of acetylated histone 4 (AC-H4), E2F1, p130, p107 and E2F4 was examined by ChIP analysis of the mouse (A) CDK1, (B) E2F1 or (C) CHEK1 promoter. For the CDK1 and E2F1 promoter, the distal E2F1 activator binding site and the proximal E2F4 repressor binding site is depicted as two blue boxes. For the CHEK1 promoter, E2F1 binding site is depicted as blue 94

108 box. The CHR binding site is depicted as the yellow box. The relative position of the PCR products used for ChIP analysis is shown by the lines with double arrows. (D) Promoter analysis of the mouse CDK1 promoter. Mutations in the distal activator E2F1 binding site, the proximal repressor E2F4 binding site and the proximal CHR binding site are illustrated. For all datasets, N = 3 independent samples per treatment group. Values represent the mean ± SEM. *significantly different than HRAS-expressing wild-type DMSO controls, P the E2f1 promoter (a gene autoregulated by E2F), in HRAS-expressing wild-type but not Pparβ/δ-null keratinocytes (Figure 2-13B). E2F4 promoter occupancy was evident on the E2F binding site and ligand activation of PPARβ/δ did not alter this occupancy (Figure 2-13B). None of these changes were observed in mock-infected keratinocytes (Figure 2-13B). Similar effects were also noted in 308 cells (Figure 2-14C). Combined, these observations suggest that ligand activation of PPARβ/δ represses HRAS-induced expression of E2F1 by increasing E2F4/p130/p107 repressor activity. Since CHEK1 is regulated by E2F1 (Carrassa et al., 2003), the effect of PPARβ/δ activation on promoter occupancy of E2F1, E2F4 and p130 was also examined. Ligand activation of PPARβ/δ caused a reduction in acetylated histone 4 and decreased promoter occupancy of E2F1 on the E2F binding site on the Chek1 promoter in HRAS-expressing wild-type but not Pparβ/δ-null keratinocytes (Figure 2-13C). No significant promoter occupancy of E2F4 or p130 to the E2F binding site was detected in HRAS-expressing keratinocytes (Figure 2-13C). To further characterize the mechanism by which ligand activation of PPARβ/δ represses CDK1 expression, mutation analysis of the CDK1 promoter was performed. Four CDK1 promoter-luciferase constructs were designed (Figure 2-13D). Ligand activation of PPARβ/δ caused repression of the wild-type CDK1 95

109 Figure Effect of ligand activation of PPARβ/δ on nuclear accumulation and promoter occupancy of p130/p107 and/or E2F in 308 cells. 308 cells were cultured to near confluency and then treated with DMSO or 1 µm GW0742 for 24 hours. (A) Western blot analysis of p107, p130 and PPARβ/δ in cytosol (C) and nuclear (N) extracts. The average ratio of nuclear to cytoplasmic protein (N/C) is shown. Promoter occupancy of AC-H4, E2F1, p130, p107 and E2F4 was examined by ChIP analysis of the mouse (B) CDK1 or (C) E2F1 promoter in 308 cells. For the CDK1 and E2F1 promoter, the distal E2F1 activator binding site and the proximal E2F4 repressor binding site is depicted as two blue boxes and the CHR binding site is depicted as the yellow box. The relative position of the PCR products used for ChIP analysis is shown by the lines with double arrows. (D) Promoter analysis of the mouse CDK1 promoter. Mutations in the proximal repressor E2F4 binding site and the proximal CHR are described in the Materials and methods. For all datasets, N = 3 independent samples per treatment group. Values represent the mean ± SEM. *significantly different than DMSO controls, P

110 promoter and the distal E2F mutant CDK1 promoter in HRAS-expressing wild-type but not Pparβ/δ-null keratinocytes (Figure 2-13D). Basal luciferase activity was significantly higher in both the proximal E2F mutant (Figure 2-13D) and the CHR mutant (data not shown), consistent with the finding that E2F4 represses CDK1 expression. However, repression of the CDK1 promoter activity was not found in response to ligand activation of PPARβ/δ with the proximal E2F mutant (Figure 2-13D) or the CHR mutant (data not shown). Similar effects were also noted in 308 cells (Figure 2-14D). These observations suggest that while E2F1 activity is dispensable, E2F4 repressor activity is indispensable for PPARβ/δ-dependent repression of CDK1 expression PPARβ/δ interacts with p107 and p130 Since nuclear translocation of PPARβ/δ in response to ligand activation in HRAS-expressing cells is concomitant with the increased nuclear accumulation of hypophosphorylated p130 and p107 (Figure 2-12A,B), this suggests that PPARβ/δ may physically interact with p130 and p107 to facilitate their translocation. It is already known that E2F4 and p130/p107 physically interact, and indeed, co-localization of p130/p107 and E2F4 was found in both wild-type and Pparβ/δ-null keratinocytes as shown by confocal microscopy (Figure 2-15A, B). In addition, co-localization of PPARβ/δ and p130/p107 was observed in HRAS-expressing wild-type, but not Pparβ/δ-null cells (Figure 2-15A, B). p107 and E2F4 were co-immunoprecipitated with PPARβ/δ (Figure 2-15C) and PPARβ/δ and E2F4 were co-immunoprecipitated with p107 in HEK293T cells 97

111 Figure PPARβ/δ binds with p107/p130. (A, B). Confocal immunofluorescence of HRAS-expressing wild-type and Pparβ/δ-null keratinocytes treated with DMSO or 1 μm GW0742 for 24 hours. Higher magnification photomicrographs are shown only for wild-type keratinocytes. Scale bar = 3 µm. Co-IP assays in HEK293T cells transiently transfected with pcmv-p107/pcmv-p130, pcmv-e2f4 and psg5-pparβ/δ (C, E), transiently transfected with FLAG-p107, pcmv-e2f4 and psg5-pparβ/δ (D), or HRAS-expressing keratinocytes (F) treated with DMSO or 1 μm GW0742 for 24 hours. (G) Interaction between endogenous p130/p107 and PPARβ/δ in HEK293T cells. 98

112 Figure PPARβ/δ binds with p107/p130 but not E2F4 and PPARβ/δ, p107/p130 and E2F4 form a large complex. (A) Co-immunoprecipitations of in vitro translated p130 and 35 S-labeled PPARβ/δ in the absence or presence of 1μM GW0742. (B, C) Co-immunoprecipitations of in vitro translated E2F4 and 35 S-labeled PPARβ/δ (B) or 35 S-labeled E2F4 and PPARβ/δ (C) in the absence or presence of 1 μm GW0742. Sequential pull down assay of HEK293T cells transiently transfected with pcmv-p107 (D) or pcmv-p130 (E), pcmv-e2f4 and pcdna-flag-pparβ/δ and treated with DMSO or 1 μm GW0742 for 24 hours. Arrows indicate p107 (D), p130 (E) and E2F4 and FLAG-PPARβ/δ respectively. (F) ChIP-re-ChIP assay on HRAS-expressing wild-type and Pparβ/δ-null keratinocytes after 24 hours of DMSO or 1 μm GW0742 treatment. Relative recruitment of PPARβ/δ, E2F4 or the complex of the two on the repressor E2F4 binding site of Cdk1 gene was quantified by qpcr. For all datasets, N = 3 independent samples per treatment group. Values represent the mean ± SEM. *significantly different than IgG controls, P (Figure 2-15D). While E2F4 and both forms of p130 were co-immunopreciptated with PPARβ/δ, hypophosphorylated p130 was preferentially pulled down (Figure 2-15E). A direct interaction between p130/p107 and PPARβ/δ was also found in HRAS-expressing primary keratinocytes (Figure 2-15F). An interaction between endogenous p130/p107 and PPARβ/δ was also found in HEK293T cells (Figure 2-15G). The findings that both p130/p107 and E2F4 were co-immunoprecipitated with PPARβ/δ suggest that either: 1) PPARβ/δ can physically bind to p130/p107 and E2F4 or 2) PPARβ/δ can bind to p130/p107 only and E2F4 was co-immunoprecipitated because E2F4 associates with p107/p130. To distinguish 99

113 between these possibilities, in vitro translated p130, PPARβ/δ and E2F4 proteins were used in a co-immunoprecipitation assay. While PPARβ/δ physically interacted with p130 (Figure 2-16A) no direct interaction between PPARβ/δ and E2F4 was observed with either E2F4 (Figure 2-16B) or PPARβ/δ pulldown (Figure 2-16C). Combined, these findings suggest that PPARβ/δ can directly interact with p130/p107 but not E2F4. The observation that both p130/p107 and E2F4 were co-immunoprecipitated with PPARβ/δ (Figure 2-15C,E) implies that p130/p107, E2F4 and PPARβ/δ may form a complex. A sequential immunoprecipitation approach was used to examine this idea. Indeed, E2F4 is detected in a complex with PPARβ/δ and p107/p130 following sequential immunoprecipitation of PPARβ/δ followed by immunoprecipitation of p107/p130 (Figure 2-16D,E), suggesting that this complex exists in a system when the three proteins are over-expressed. To determine if this complex is found on the promoter of the Cdk1 gene, a ChIP-re-ChIP assay was performed using a crosslinker (DTBP) that allows detection of proteins that are not directly bound to chromatin. With this approach, promoter occupancy of PPARβ/δ was detected at the same site in HRAS-expressing wild-type cells and increased in the presence of GW0742 (Figure 2-16F) but PPARβ/δ is not detected on the E2F4 repressor sites using formaldehyde as the crosslinker (data not shown). Enriched promoter occupancy of E2F4 and PPARβ/δ was found after sequential PPARβ/δ and E2F4 pull-down only in wild-type cells (Figure 2-16F). These data suggest that PPARβ/δ, E2F4 and p130/p107 may form a complex on the Cdk1 promoter. Whether this may occur for other E2F target genes remains to 100

114 Figure Ligand activation PPARβ/δ attenuates phosphorylation of p130 by CDKs. (A) Commassie staining of affinity purified recombinant GST-p107 and GST-p130 protein isolated from E. coli. In vitro kinase assay was performed and the ratio of the phosphorylated p130 relative to total p130 (B) or phosphorylated p107 relative to total p107 (C) is shown below each band. (D) Co-IP assays were performed with HRAS-expressing keratinocytes treated with DMSO or 1μM GW0742 for 24 hours. Arrows indicate CDK2, CDK4, E2F4 or p130. The amount of CDK2, CDK4 and E2F4 pulled down was quantified by normalizing to the amount of p130 pulled down. be determined. PPARβ/δ may preferentially interact with hypophosphorylated p130 (Figure 2-15E), suggesting that the binding of PPARβ/δ to p130 may protect p130 from phosphorylation. Since p130 can be phosphorylated by CDK4/CYCLIN D1 complex (Hansen et al., 2001), an in vitro kinase assay was performed to examine the hypothesis that PPARβ/δ preferentially interacts with hypophosphorylated p130. Purifity of GST-p130 and GST-p107 recombinant protein was confirmed by Commassie staining (Figure 2-17A). The addition of both PPARβ/δ and GW0742 decreased the phosphorylation of p130 by 33% percent (Figure 2-17B). The 101

115 decreased phosphorylation of p130 was not due to competition between PPARβ/δ and p130 for CDK4/CYCLIN D1 because PPARβ/δ was not phosphorylated by CDK4/CYCLIN D1 (Figure 2-17B). To determine whether the observed decrease of phosphorylation of p130 by ligand activation of PPARβ/δ was due to the decreased binding of CDK4/CYCLIN D1 complex to p130, the interaction of p130 and CDK4 was examined. Ligand activation of PPARβ/δ decreased the association between p130 and CDK4 in HRAS-expressing cells (Figure 2-17D). In addition, the interaction between p130 and CDK2, which can also phosphorylate p130 (Cheng et al., 2000), was also decreased by ligand activation of PPARβ/δ (Figure 2-17D). No significant change in the phosphorylation of p107 by CDK4/CYCLIN D1 in the presence of PPARβ/δ and/or GW0742 was observed (Figure 2-17C) Ligand activation of PPARβ/δ attenuates mitosis in vivo To determine if the changes found in HRAS-expressing cells in response to ligand activation of PPARβ/δ are also observed in vivo, the mitotic index and expression of Hras was examined in skin tumors obtained from a two-stage bioassay (initiation with DMBA, promotion with TPA). Ligand activation of PPARβ/δ caused a decrease in mitotic index in skin tumors from wild-type, but not Pparβ/δ-null mice (Figure 2-18A-C). In addition, the mitotic index in skin tumors from Pparβ/δ-null mice was higher as compared to wild-type mice (Figure 2-18A-C). Consistent with the hypothesis that PPARβ/δ-dependent inhibition of mitosis causes selection against cells expressing higher levels of HRAS, expression of 102

116 Figure Ligand activation of PPARβ/δ selects against higher HRAS-expressing chemically-induced skin tumors by inhibiting mitosis. (A) Representative photomicrographs of skin tumors in wild-type or Pparβ/δ-null mouse skin treated with or without GW0742. The mitotic index was calculated from a minimum of 1000 cells per sample. Quantification of mitotic index from samples examined in (A) excluding cells in the G2/M boundary (B) and including cells in the G2/M boundary (C). For B and C, representative skin tumors were used (N = 5-8). (D) RNA was isolated from skin tumors and Hras mrna 103

117 expression was determined by qpcr and normalized to 18s RNA. For all datasets, N = 6-10 independent samples. Values represent the mean ± SEM. *significantly different than wild-type controls, P Scale bar = 100 µm. Western blot analysis of (E) CDK1, CHEK1, E2F1 and HRAS or (F) p130, p107, E2F4 and PPARβ/δ in skin and skin tumors. Expression levels of proteins were normalized to β-actin and are presented as (E) the fold change relative to control DMSO, or as (F) the average ratio of nuclear to cytoplasmic protein (N/C). Downward arrow in (E) indicates the same wild-type control sample used for comparison. (G) Co-IP assay showing an interaction between PPARβ/δ and p107/p130 in cell lysates from skin tumors treated with GW0742. Hras mrna was lower in skin tumors from wild-type mice treated with GW0742, an effect not found in Pparβ/δ-null mice (Figure 2-18D). In addition, ligand activation of PPARβ/δ also decreased the level of proteins that promote mitosis including CDK1, CHEK1 and E2F1 in skin tumors from wild-type, but not in Pparβ/δ-null mice (Figure 2-18E). Expression of HRAS was also reduced by ligand activation of PPARβ/δ in wild-type mouse skin tumors but not in Pparβ/δ-null mouse skin tumors (Figure 2-18E). Consistent with results observed in HRAS-expressing primary keratinocytes and 308 cells, ligand activation of PPARβ/δ increased the nuclear to cytosol ratio of p130 (hypophosphorylated), p107, E2F4 and PPARβ/δ in skin tumors but not in adjacent non-transformed skin (Figure 2-18F). There was also an increase in nuclear accumulation of phosphorylated p130 in skin tumors following ligand treatment (Figure 2-18F). There are at least two possibilities why the two forms of p130 increase when PPARβ/δ is activated. First, even though PPARβ/δ preferentially interacts with hypo-p130, PPARβ/δ can also interact with phosphorylated p130 (Figure 2-15E, F). Thus, when PPARβ/δ is activated, nuclear translocation of PPARβ/δ may lead to an increase in both hypo and phosphorylated p130. The second possibility is 104

118 that even though ligand activated PPARβ/δ decreases phosphorylation of p130 (Figure 2-17B), it does not completely prevent p130 from being phosphorylated by CDKs. Thus, nuclear hypo-p130 may be phosphorylated by CDK2/CDK4 complex that are present in the nucleus and this may account for the observed increased both forms of p130 in the nucleus when PPARβ/δ is activated. Association between PPARβ/δ and p107 and hypo-phosphorylated p130 was also detected in wild-type skin tumors treated with GW0742 (Figure 2-18G). These findings suggest that ligand activation of PPARβ/δ also attenuates mitosis in chemically-induced skin tumors with an HRAS mutation through cross-talk with E2F signaling Enhanced sensitivity to pharmacological inhibition of mitosis in HRAS-expressing cells by ligand activation of PPARβ/δ Other therapeutics can effectively inhibit growth of transformed cells by blocking progression at the M phase of the cell cycle including RO-3306 (CDK1 inhibitor), paclitaxel (microtubule stabilizer), nocodazole (microtubule de-stabilizer) and SB (CHEK1 inhibitor). Since ligand activation of PPARβ/δ with GW0742 also causes G2/M arrest in HRAS-expressing keratinocytes, the effect of combining GW0742 with other mitosis inhibitors on cell proliferation was examined. RO-3306 increased the percentage of cells in the sub G1 and G2/M phase of the cell cycle in HRAS-expressing keratinocytes and this effect was markedly increased by co-treatment with GW0742; an effect not found in similarly treated HRAS-expressing Pparβ/δ-null keratinocytes (Figure 2-19A, B). These 105

119 Figure Ligand activation of PPARβ/δ leads to hypersensitivity to pharmacological inhibition of mitosis in HRAS-expressing keratinocytes. (A-D) HRAS-expressing wild-type or Pparβ/δ-null keratinocytes were treated with or without 1 µm GW0742 for 2 d and then treated with either 1 µm GW0742, 10 µm RO-3306, or 10 µm RO-3306 and 1 µm GW0742 for 24 h. (A) Representative photomicrographs after 24 h of co-treatment; a significant increase in cell death was observed in HRAS-expressing wild-type keratinocytes co-treated with GW0742 and RO-3306 but not Pparβ/δ-null keratinocytes. (B) Distribution of cells in sub-g1 and G2/M phase of the cell cycle. (C) The percentage of viable cells was normalized to its own control in the absence of RO (D). Flow cytometric analysis using an anti-hras antibody was performed and Pearson s second skewness coefficient of HRAS intensity was calculated. (E) HRAS-expressing keratinocytes were cultured in medium with or without 1 µm GW0742 or treated with 20 nm paclitaxel or 100 ng/ml nocodazole with or without 1 µm GW0742 for 40 h. Distribution of cells in the G2/M phase of the cell cycle was determined. (F, G) HRAS-expressing wild-type or Pparβ/δ-null keratinocytes were treated with or without 1 µm GW0742 for 2 d and then treated with either 1 106

120 µm GW0742, 10 µm SB218078, or 10 µm SB and 1 µm GW0742 for 24 h. (F) Representative photomicrographs after 24 h of co-treatment; a significant increase in cell death was observed in HRAS-expressing wild-type keratinocytes co-treated with GW0742 and SB but not Pparβ/δ-null keratinocytes. (G) The percentage of viable cells was normalized to its own control in the absence of SB For all datasets, N = 3 independent samples per treatment group. Values represent the mean ± SEM. *significantly different than HRAS-expressing wild-type DMSO controls, P changes were consistent with enhanced PPARβ/δ-dependent inhibition of cell proliferation observed in HRAS-expressing wild-type keratinocytes co-treated with RO-3306 and GW0742 (Figure 2-19C). Similar to the effect of ligand activation of PPARβ/δ, treatment with RO-3306 caused selection against cells expressing higher levels of HRAS in wild-type keratinocytes (Figure 2-19D). Co-treatment of RO-3306 with GW0742 enhanced this selection (Figure 2-19D). Paclitaxel also increased the percentage of cells in the G2/M phase of the cell cycle in HRAS-expressing keratinocytes and this effect was increased by co-treatment with GW0742; an effect not found in HRAS-expressing Pparβ/δ-null keratinocytes (Figure 2-19E). In contrast, co-treatment of GW0742 with nocodazole did not increase the effects induced by nocodazole alone (Figure 2-19E). Similar enhanced inhibition of cell proliferation was also found following co-treatment with GW0742 and RO-3306 or paclitaxel in 308 mouse keratinocytes (data not shown). Enhanced PPARβ/δ-dependent inhibition of cell proliferation was also observed in HRAS-expressing wild-type cells co-treated with the CHEK1 inhibitor SB and GW0742 (Figure 2-19F, G). 107

121 2.5 Discussion These studies are the first to demonstrate PPARβ/δ-dependent inhibition of cell proliferation in HRAS-expressing cells by increasing G2/M arrest. This is consistent with previous work that showed inhibition of skin carcinogenesis and inhibition of proliferation in keratinocyte cell lines with Hras mutations (Bility et al., 2008; Bility et al., 2010). The current studies revealed that induction of G2/M arrest caused by ligand activation of PPARβ/δ specifically targets cells with higher expression of HRAS. That the PPARβ/δ-dependent induction of G2/M arrest causes selection against cells with higher expression of HRAS is consistent with results showing that cells treated with chemicals that induce G2/M arrest also cause selection against cells expressing higher levels of HRAS. Interestingly, human cancer cell lines expressing oncogenic RAS are more sensitive to mitotic perturbations as compared to normal cells (Luo et al., 2009), an observation also noted in the present studies. For example, as HRAS activity increased in keratinocytes (by viral transduction) or 308 cells (with an activating mutation in HRAS), the efficacy of GW0742 to induce G2/M arrest increased. Moreover, ligand activation of PPARβ/δ inhibits mitosis in skin tumors and this phenotype is also associated with reduced expression of HRAS in these tumors. It was also shown that cells expressing oncogenic RAS exhibit a disadvantage for cell proliferation following knockdown of many mitotic genes (Luo et al., 2009). This is important to note because PPARβ/δ-dependent repression of many of these mitosis related genes was also observed in HRAS-expressing keratinocytes, 308 cells and skin tumors in the present study. 108

122 Of the mitosis-related genes that were repressed by ligand activation of PPARβ/δ in HRAS-expressing cells, Cdk1 and Chek1 are of great interest. While some of these changes were relatively modest, this could be due to the presence of an endogenous high affinity agonist that may prevent more robust alterations in expression. An active CYCLIN B1-CDK1 complex is a trigger to enter mitosis whereas depletion of CYCLIN B1-CDK1 can cause a block in mitosis, concomitant with repeated rounds of S phase, leading to cells with polyploidies in both fission yeast and human cells (Correa-Bordes and Nurse, 1995; Hayles et al., 1994; Itzhaki et al., 1997; Moreno and Nurse, 1994). CHEK1 is required for spindle checkpoint function (specifically at the tension sensing branch of the checkpoint) and Chek1 / cells can exit mitosis in the presence of paclitaxel and undergo endoreduplication, leading to polyploidies (Zachos et al., 2007). Ligand activation of PPARβ/δ decreased expression of CDK1 and CHEK1 in HRAS-expressing cells and the observed phenotype, including delayed entry into mitosis, retarded exit from mitosis, and increased polyploidy cells is similar to the phenotype of Cdk1-null and Chek1-null cells. Combined, these observations suggest that the PPARβ/δ-dependent decrease in expression of CDK1 and CHEK1 alone in HRAS-expressing cells may largely underlie the observed mitosis block following ligand activation of PPARβ/δ. Inhibition of cell cycle kinetics induced by ligand activation of PPARβ/δ could be due in part to direct regulation of target genes by PPARβ/δ, which was not examined in the present study. However, results from these studies also establish that PPARβ/δ-dependent inhibition of mitosis in cells with an activating Hras 109

123 Figure Mechanism of PPARβ/δ mediated inhibition of mitosis in HRAS expressing cells. PPARβ/δ chaperones p130 into the nucleus where it represses E2F target genes causing inhibition of mitosis. mutation can also inhibit cell cycle progression and is mediated by a mechanism (Figure 2-20) that involves: 1) PPARβ/δ directly binding with p107/p130 proteins; 2) translocation of PPARβ/δ to the nucleus in response to ligand activation leading to increased nuclear hypophosphorylated p130 and p107; 3) ligand bound-pparβ/δ maintaining p130 in a hypophosphorylated state; and 4) heightened nuclear p107/p130 causing increased recruitment of p130/p107/e2f4 complex to the promoters of mitosis related genes and inhibition of their transcription; genes with repressor E2F4 binding sites (such as Cdk1 and E2f1) 110

124 that are repressed directly by this complex. Because of this PPARβ/δ-dependent down-regulation of E2F1 expression, decreased E2F1 recruitment to promoters of other genes preferentially regulated by activator E2Fs (such as Chek1) are secondarily influenced by this regulation. Thus, it is not surprising that expression of CHEK1 is also down-regulated by ligand activation of PPARβ/δ. Alternatively, it remains possible that the PPARβ/δ/p130/p107/E2F4 complexes could exhibit differential affinity for binding sites on chromatin, or lead to differences in recruitment of transcriptional co-repressors. Further studies are needed to examine these ideas. Since cells with RAS mutations are more sensitive to mitotic perturbations compared to normal cells, the present studies focused more on the regulation of mitosis genes. However, it is also noteworthy that expression of many E2F target genes involved in DNA replication and DNA repair was also reduced by ligand activation of PPARβ/δ. This change in gene expression is reflected by the decrease in cells undergoing S phase in HRAS-expressing keratinocytes following ligand activation of PPARβ/δ. The interaction between p107/p130 and PPARβ/δ is independent of HRAS. Moreover, PPARβ/δ preferentially binds to the hypophosphorylated form of p130 based on data from co-immunoprecipitations. This is also supported by results showing more prominent co-localization of p130 and PPARβ/δ in the cytosol of cells, since hypo-phosphorylated p130 was found primarily in the cytoplasm. In the presence of ligand, PPARβ/δ may inhibit p130 from being phosphorylated by CDK4. This suggests that when p130 is shuttled to the nucleus via random nuclear translocation of PPARβ/δ under normal conditions, p130 will be 111

125 phosphorylated by CDK4/CYCLIN D1 complex present in the nucleus, thus losing its repressor activity. This also suggests that ligand activation of PPARβ/δ is essential for repression of mitosis genes because it maintains p130 in a hypophosphorylated state and chaperones hypophosphorylated p130 into the nucleus in cells with activated HRAS signaling. Nuclear translocation of PPARβ/δ in HRAS-expressing cells following ligand activation is central to the inhibition of mitosis genes. Nuclear translocation of PPARβ/δ and increased nuclear p107 and p130 in normal keratinocytes are typically not observed. In contrast, increased nuclear translocation of PPARβ/δ and concurrent increase of p107 and p130 in HRAS-expressing keratinocytes and skin tumors illustrates the essential nature of nuclear translocation of PPARβ/δ following ligand activation in the presence of activated HRAS signaling. The increase in both cytosolic and nuclear PPARβ/δ following ligand activation in control keratinocytes is most likely due to the stabilization of the receptor rather than an increase of protein synthesis and nuclear translocation for the following reasons. First, no increase in PPARβ/δ mrna was observed following ligand activation. Second, ligand activation of PPARβ/δ is known to prevent its ubiquitin-mediated degradation, thus increasing its half-life (Genini and Catapano, 2007). However, the mechanism of nuclear translocation of PPARβ/δ in HRAS-expressing keratinocytes, skin tumors and confluent 308 cells following ligand activation remains unclear. It is possible that increased HRAS activity may activate downstream kinases and alter the phosphorylation status of PPARβ/δ, leading to its nuclear translocation. 112

126 Increasing G2/M arrest of cells expressing high levels of HRAS can be achieved by ligand activation of PPARβ/δ, and co-treatment of a PPARβ/δ ligand with various mitosis inhibitors enhances the efficacy of increasing a G2/M arrest. This supports the hypothesis that combining ligand activation of PPARβ/δ with mitosis inhibitors may be a feasible approach for treating tumors that express higher levels of RAS. Indeed, oncogenic RAS signaling is increased in a number of human cancers including lung, colon, pancreas, and melanoma (Schubbert et al., 2007). While the role of PPARβ/δ in some cancers remains controversial (reviewed in (Peters et al., 2011a; Peters and Gonzalez, 2009; Peters et al., 2008; Peters et al., 2012; Peters et al., 2011b)), the body of evidence suggesting that PPARβ/δ protects against cancer is increasing. For example, a recent compelling study demonstrated that colorectal cancer patients with relatively low expression of PPARβ/δ in the primary tumor were ~4X as likely to die from this disease as compared to patients with relatively higher expression of PPARβ/δ in their primary tumors (Yang et al., 2011b). It is also not disputed that ligand activation of PPARβ/δ inhibits chemically-induced skin carcinogenesis (Bility et al., 2008; Bility et al., 2010; Kim et al., 2006; Zhu et al., 2011). Moreover, preclinical and clinical studies have also shown that ligand activation of PPARβ/δ inhibits or prevents metabolic syndrome, obesity, dyslipidemias, glucose intolerance and chronic inflammation; characteristics that are positively associated with cancer (Pais et al., 2009; Prizment et al., 2010; Terzic et al., 2010; Tsugane and Inoue, 2010; Wolin et al., 2011). Since targeting single molecules for chemoprevention and chemotherapy has not proven highly effective (Hanahan and Weinberg, 2011) due 113

127 in part to the genetic heterogeneity associated with diseases (Schadt, 2009), targeting PPARβ/δ in conjunction with mitosis inhibitors could become a suitable option for development of new multi-target strategies for inhibiting RAS-dependent tumorigenesis. 114

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136 Chapter 3 PPARβ/δ promotes HRAS-induced senescence and tumor suppression by regulating p-erk, p-akt and ER Stress 3.1. Abstract PPARβ/δ inhibits skin tumorigenesis through mechanisms that may be dependent on HRAS signaling. The present study examined the hypothesis that PPARβ/δ promotes HRAS-induced senescence thereby suppressing tumorigenesis. PPARβ/δ transcriptionally upregulates Rasgrp1 and suppresses Ilk causing increased p-erk and decreased p-akt that promote HRAS-induced senescence. PPARβ/δ also promotes senescence through attenuation of HRAS-induced endoplasmic reticulum (ER) stress by repressing p-akt-mtor signaling, and repressing ER stress-associated unfolded protein response (UPR). Decreased UPR increases senescence, whereas increased ER stress and UPR decrease HRAS-induced cellular senescence. Further, these studies demonstrate a novel positive feedback loop between p-akt, ER stress and UPR. An acute increase of ER stress is sufficient to establish the positive loop, maintaining higher UPR and p-akt activity, collectively causing evasion of senescence and malignant conversion. Moreover, increased PPARβ/δ expression correlates with increased senescence and decreased ER stress correlates with increased senescence in both mouse and human tumors. These results are the first to demonstrate that PPARβ/δ promotes senescence and inhibits tumorigenesis. These studies also identify a new pro-tumorigenic role of ER stress facilitated by attenuation of senescence. 123

137 3.2 Introduction The role of peroxisome proliferator-activated receptor-β/δ (PPARβ/δ) in some cancers remains controversial, but there is strong evidence that PPARβ/δ attenuates non-melanoma skin cancer (reviewed in (Peters et al., 2012)). Pparβ/δ-null mice exhibit exacerbated skin tumorigenesis and ligand activation of PPARβ/δ inhibits chemically-induced skin tumorigenesis; likely mediated by PPARβ/δ-dependent induction of terminal differentiation and inhibition of cell proliferation and mitosis (Bility et al., 2008; Bility et al., 2010; Kim et al., 2004a; Zhu et al., 2010). Mutations in the Harvey sarcoma ras virus gene (Hras) is found in over 90% of 7,12-dimethylbenz[a]anthracene (DMBA) initiated skin tumors (Quintanilla et al., 1986) and increased activity associated with mutant HRAS is one mechanism that can cause cancer (Schubbert et al., 2007). An activated mutant HRAS (Hras-V12) triggers cellular senescence (irreversible cell cycle arrest), concomitant with increased expression of p16 and p53 tumor suppressors (Serrano et al., 1997). Oncogene-induced senescence has also been observed for a number of other mutations in other genes including KRAS, BRAF, PTEN and NF1 (Courtois-Cox et al., 2008). It is widely believed that oncogene-induced senescence serves as a self-defense mechanism to suppress tumor development by preventing the progression of benign lesions to malignancies in the absence of additional co-operating mutations (Courtois-Cox et al., 2008). Further, it was also recently suggested that the extent of endoplasmic reticulum (ER) stress induced by mutant HRAS contributes to the mechanism underlying cellular senescence (Denoyelle et al., 2006). The idea that 124

138 that an interaction exists between ER stress and cellular senescence in the presence of mutant HRAS has not been extensively examined to date. Thus, the present study examined the mechanisms by which PPARβ/δ attenuates HRAS-dependent skin tumorigenesis, with an emphasis on cellular senescence and ER stress. 3.3 Materials and methods Plasmids and Vectors pbabe-puro-mek-dd as previously described (Boehm et al., 2007) was obtained from Addgene (plasmid #15268). pbabe-puro-pten was obtained by cloning mouse PTEN from mouse keratinocytes mrna and cloned into BamHI and EcoRI sites of pbabe-puro vector. pbabe-puro-mouse Rasgrp1 was obtained by PCR-amplifying mouse Rasgrp1 ORF from mouse-rasgrp1 shuttle clone vector (GC-Mm04880) (GeneCopoeia, Rockville, MD) and cloning into BamHI sites of pbabe-puro vector. Non-target control shrna and mouse shrna against Ilk, Atf4 and Xbp1 were purchased from Mission shrna (Sigma-Aldrich, St.Louis, MO). The shrna catalogue number and sequences are as follows: non-target control shrna: (SHC002): CCGGCAACAAGATGAAGAGCACCAACTCGAGTTGGTGCTCTTCATCTTGTT GTTTTT, mouse Ilk shrna1 (TRCN ): CCGGGCACGGATTAATGTGATGAATCTCGAGATTCATCACATTAATCCGTGC 125

139 TTTTT, mouse Ilk shrna2 (TRCN ): CCGGCCTGAACAAACACTCCGGTATCTCGAGATACCGGAGTGTTTGTTCAG GTTTTT, mouse Atf4 shrna (TRCN ): CCGGCGGACAAAGATACCTTCGAGTCTCGAGACTCGAAGGTATCTTTGTCC GTTTTT, mouse Xbp-1 shrna (TRCN ): CCGGCCATTAATGAACTCATTCGTTCTCGAGAACGAATGAGTTCATTAATGG TTTTT Cell culture Primary keratinocytes from newborn wild-type and Pparβ/δ-null mice were prepared and cultured as previously described (Dlugosz et al., 1995). Keratinocytes were infected with the Hras retrovirus for two days at an estimated MOI of 3 as previously described (Roop et al., 1986). HaCat cell line was maintained in DMEM medium as previously described (Borland et al., 2008). 1 μm GW0742, 10 μm LY and 10 μm PD98059 were used throughout the manuscript unless otherwise indicated. 3μg/ml Anti-BiP antbody (N-20) (Santa Cruz technology, Santa Cruz, CA) was used to immunoneutralize cell-surface BiP as this concentration was previously shown to be able to completely block Cripto binding (Kelber et al., 2009). 126

140 3.3.3 Virus production and keratinocytes infection The Hras retrovirus was generated from ψ2 producer cells as described previously (Roop et al., 1986). The virus titer was determined to be between 1 2 x 10 7 transforming units/ml using an NIH-3T3 focus-forming assay. To obtain shrna lentivirus, 24 μg shrna vector, 16 μg PAX2 (or HIV-gag/pol) and 8 μg MD2G (or VSV-G) packaging plasmids were transiently tranfected into HEK293T cells. 24 hours later, virus-containing supernatant were filtered through 0.45 μm filter and virus were precipitated with PEG-it virus precipitation solution (System Biosciences, Mountain View, CA) and resuspended with low calcium medium. Keratinocytes were first infected with Hras retrovirus for 24 hours before infected with shrna retrovirus with an approximate M.O.I of 5 for 24 hours. After 24 hours of recovery in normal low calcium medium, 1 μg /μl puromycin were added to the medium to select for puromycin resistant cells. To produce virus from pbabe retroviral vectors, we utilized ψ2 producer cells (which already has a packaging plasmid stably integrated into the genome) to produce a mixture of Hras and pbabe-based retrovirus. pbabe empty vector, pbabe-puro-mek-dd, pbabe-puro-pten or pbabe-puro-mouse Rasgrp1 was electroporated into ψ2 producer cells using 4D-nucleofactor apparatus (Lonza, Walkersville, MD). Stably integrated cells were selected after of culture in medium containing 1 μg /μl puromycin. Viruses were collected and keratinocytes were infected with virus mixture and puromycin resistant keratinocytes were selected as described above. 127

141 3.3.4 Complete carcinogenesis bioassay Female C57BL/6 mice, in the resting phase of the hair cycle (6~8 weeks of age) on either wild-type or Pparβ/δ-null background were initiated with 50 μg of 7, 12-dimethylbenz[a]anthracene (DMBA) dissolved in 200 μl acetone. One week after initiation, mice were treated topically with 50 μg of DMBA weekly. Meanwhile, mice were also topically treated with acetone or 5 μm GW0742 twice a week for 30 weeks. After 30 weeks, mice were euthanized by overexposure to carbon dioxide. Tumor samples were either fixed or snap frozen in liquid nitrogen for future analysis. Fixed tumor samples were embedded in paraffin, sectioned and stained with hematoxylin and eosin (H&E) and scored for benign or malignant pathology by pathologists In vitro malignant transformation assay Primary keratinocytes after pre-attachment treatment to remove fibroblasts were plated at high density and infected with v-hras1 retrovirus on day 2 of culture. Infected keratinocytes were cultured for 6 days in Eagle's Minimal Essential Medium containing 0.05 mm calcium and 8% FBS before switched to 0.5 mm calcium medium and treated with indicated chemicals for 4 weeks. Colonies of keratinocytes resistent to calcium-induced differentiation were identified by staining the cells with 0.35% rhodamine/10% formalin. Rhodamine stained-calcium resistant colonies were counted with a dissecting microscope to eliminate contaminated fibrloblasts and colony diameter was measured using a micrometer. In a separate experiemnt, six 0.5 mm calcium-resistant 128

142 HRAS-expressing Pparβ/δ-null keratinocytes monoclones were successfully isolated Clonogenic assay with ER stress inducers Primary keratinocytes after pre-attachment treatment to remove fibroblasts were plated at high density and infected with v-hras1 retrovirus for 48 hours. For short term treatment, infected keratinocytes were treated with DMSO, 10μM PD98059, 1 nm thapsigargin, 2.5 nm thapsigargin or a combination of 10μM PD98059 and 1 nm thapsigargin for 4 days in Eagle's Minimal Essential Medium containing 0.05 mm calcium and 8% FBS. Chemicals were removed after 4days of treatment and cells were either maintained in low calcium medium or switched to 0.5 mm calcium medium for 4 weeks. For continued treatment, cells were maintained in medium containing 1 nm thapsigargin throughout culture. Colonies were first stained with anti-keratin 5 (COVANCE, Emeryville, CA) antibody to eliminate contaminated fibroblasts and then identified by staining the cells with 0.35% rhodamine/10% formalin Cell proliferation assays Keratinocytes were seeded in 96 well plates, infected with Hras retrovirus and after seventy-two hours of treatment with LY294002, PD98059 or tunicamycin, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) in PBS was added to each well at the final concentration of 0.5 mg/ml and cells were 129

143 incubated at 37 C for 1 hour. The media was removed to quantify optical density with a spectrophotometer at 560 nm after dissolving MTT in DMSO Senescence-associated β-galactosidase (SA-β-gal) assay Cells were washed twice with phosphate-buffered saline (PBS; ph 7.2), fixed with 0.5% glutaraldehyde in PBS and washed in PBS supplemented with 1 mm MgCl 2. Cells were stained at 37 C in X-Gal solution (1 mg/ml X-Gal, 0.12 mm K 3 Fe[CN] 6, 0.12 mm K 4 Fe[CN] 6, 1 mm MgCl 2 in PBS at ph 6.0). The staining was performed for 24h. For tumor samples, samples were embedded in OCT, cryosectioned and stained as previously described with the exception that the staining was performed for 4 6 h to minimize the background signal Flow cytometry analysis Cells were stained with bromodeoxyuridine (BrdU) and propidium iodide (PI) and analyzed for cell cycle progression as previously described in chapter two. For analysis of DNA content, cells were trypsinized and washed with phosphate buffered saline (PBS) once before overnight fixation in 70% ethanol. Cells were then washed with PBS and stained with PI. Approximately 10,000 cells/sample were analyzed by flow cytometry to detect PI using a FC500 flow cytometer (Beckman Coulter, Miami Lakes, FL). The percentage of cells at each phase of the cell cycle ± S.D. was determined with FCS Express software. For anti-hras staining analysis, cells were trypsinized and washed with PBS once before 130

144 fixation in 2% formaldehyde in PBS for 15 minutes at room temperature followed by permeabilization with 100% methanol for 10 minutes at -20 C. Cells were washed with PBS and incubated overnight with an anti-hras antibody (C20, Santa Cruz Biotechnology, Santa Cruz, CA) followed by incubation with an Alexa-488 conjugated secondary antibody (Cell Signaling, Beverly, MA) for one hour at room temperature in the dark. Approximately 5,000 cells/sample were analyzed by flow cytometry using a Coulter XL-MCL (Beckman Coulter, Miami Lakes, FL). Pearson s second skewness coefficient, defined as 3 (mean relative HRAS intensity median relative HRAS intensity)/ standard deviation of relative HRAS intensity was calculated to determine the relative distribution of cells with varying levels of HRAS Immunofluorescence analysis For in vitro cell culture, HRAS-expressing wild-type and Pparβ/δ-null keratinocytes were cultured in chamber slides to ensure complete attachment. Cells were fixed in 2% formaldehyde in PBS for 15 minutes at room temperature followed by permeabilization with 100% methanol for 10 minutes at -20 C. Cells were then washed with PBS, blocked with 10% normal serum of the species of the secondary antibody used for 30 minutes at room temperature and incubated overnight with primary antibodies at 4 C followed by incubation with Alexa-488, Alexa-568 or Alexa-647 conjugated secondary antibodies (Life Technologies, Grand Island, NY) for 1 hour at room temperature in the dark. Alexa-conjugated primary antibodies were incubated together with secondary antibody. Cells were 131

145 then washed with PBS before incubation in 1 μg/ml Hoechst for 10 minutes at room temperature to counter-stain DNA. For ER staining, ER-Tracker Blue-White DPX was used as indicated by the provider (Molecular Probes, Carlsbad, CA). For in vivo tumor samples, frozen sections were fixed with ice-cold acetone at -20 C for 10 minutes. Sections were air dried at room temperature for 20 minutes, re-hydrated with PBS for 15 minutes and subject to block and incubation with antibody as described above. The following antibodies were used: anti-p16 (F-12), anti-grp78 (N-20), anti-dcr2 (D-15), anti-xbp-1 (F-4) (which can recognize both spliced and unspliced XBP-1), anti-atf4 (C-19), Alexa-647 conjugated anti-p27 (F-8) (Santa Cruz technology, Santa Cruz, CA), anti-p-akt (S473) (#4058) (Cell Signaling, Beverly, MA), Alexa-555 conjugated anti-e-cadherin, Alexa-555 conjugated anti-ki-67 (BD Biosciences Pharmingen, San Diego, CA), Representative photomicrographs were obtained with an Olympus Fluoview 1000 confocal microscope. Photomicrographs were acquired with FV10-ASW2.0 VIEWER software (Olympus) and mean intensity of fluorescence emission (both cytosol and nuclear level) of at least 1000 individual cells was quantified using Cell Profiler software (Broad Institute). Intensities calculated from photomicrographs taken from different tumor specimens (at least four tumors each genotype) were normalized so that the highest and lowest intensity from different photomicrographs are equal. 132

146 Protein synthesis assay The Click-iT AHA Alexa Fluro 488 protein synthesis HCS kit (Life Technologies, Grand Island, NY) was used to measure protein synthesis in vitro as recommended by manufacturer s protocol. Briefly, HRAS-expressing keratinocytes grown in chamber slides were treated with different chemicals for 3 days before pulsed with 50μM L-azidohomoalanine in methionine-free medium for 1 hour. Cells were fixed with 2% formaldehyde and permeabilized by 0.5% Triton X-100 and undergo ligation reaction for 30 minutes in the dark. Pictures were taken by a material scope (OLYMPUS BX61) and mean intensity of Alexa-488 signal from at least 1000 cells was measured using Cell Profiler software (Broad Institute) Reactive Oxygen Species (ROS) detection assay Image-iT live green reactive oxygen species detection kit (Life Technologies, Grand Island, NY) was used to measure level of ROS in HRAS-expressing keratinocytes. Wild-type HRAS-expressing keratinocytes were treated with 2.5 nm thapsigargin or 25ng/μl tunicamycin for 4 days. After treatment, cells were washed with Hank s balanced salt solutions and incubated with 25 μm 5-(and-6)-carboxy-2',7'-dichlorodihydrofluorescein diacetate (carboxy-h 2 DCFDA) for 30 minutes at 37 C. Cells were washed three times with Hank s balanced salt solutions before subject to imaging using fluorescence microscope. Mean intensity of fluorescence signal from at least 1000 cells was measured using Cell Profiler software (Broad Institute). 133

147 RNA isolation, quantitative real-time PCR (qpcr) and RT-PCR-based detection of XBP-1 splicing Total RNA was isolated from cell lines using TRIZOL reagent (Invitrogen, Carlsbad, CA). Reverse transcription and qpcr was performed as previously described (Kim et al., 2004a). The relative level of mrna was normalized to that of glyceraldehyde 3-phosphate dehydrogenase (Gapdh) or 18s RNA levels. For detection of XBP-1 spliced (s: 162 bp) and unspliced form (us: 188 bp), forward primer (5 - AAGAACACGCTTGGGAATGGACAC- 3 ) and reverse primer (5 - ACAGTGTCAGAGTCCATGGGAAGA 3 ) were used. Primers of other genes for qpcr are available on request Quantitative western blot analysis For total cell lysates, cells were lysed with RIPA buffer supplemented with 30 mm sodium fluoride, 40 mm glycerophosphate, 10 mm sodium pyrophosphate, 2 mm sodium orthovanadate and a protease inhibitor cocktail. The supernatant after centrifugation was used for immunoprecipitations or western blot analysis. Western blot analysis using radioactive detection methods was performed as previously described (29). The primary antibodies used were: anti-phospho-retinoblastoma (Ser780) (#9307), anti-phospho-mek1/2 (Ser217/221) (#9121), anti-mek1/2 (#9122), anti-phospho-erk1/2 (Thr202/Tyr204) (#9101), anti-erk1/2 (#4695), anti-phospho-akt (Ser473) (#4058), anti-phospho-akt (Thr308) (#9275), anti-akt (#9272), anti-p27kip1 (#2552), anti-pten (#9552), anti-bip (#3177), anti-calnexin (#2679), 134

148 anti-ire1α (#3294), anti-phospho-perk (Thr980) (#3179), anti-perk (#5683), anti-phospho-s6rp (Ser235/236) (#2211), anti-s6rp (#2317), anti-phospho-4e-bp1 (Thr37/46) (#2855), anti-4e-bp1 (#9644), anti-phospho-p70 S6K (Thr389) (#9205), anti-parp (#9542) (Cell Signaling, Beverly, MA), anti-hras (C-20), anti-retinoblastoma (C-15), anti-p16 (M-156), anti-p53 (FL-393), anti-p21 (C-19), anti-dcr2 (D-15), anti-rasgap120 (171), anti-rasgrp1 (H-120), anti-dusp1 (H-66), anti-xbp1 (M-186), anti-atf4 (C-19), anti-atf6 (H-280), anti-bax (I-19), anti-pcna (PC10) (Santa Cruz Biotechnology, Santa Cruz, CA), anti-lactic dehydrogenase (LDH; Jackson Immunoresearch, West Grove, PA), anti-β-actin (Rockland, Gilbertsville, PA), anti-bad, anti-erk1/2 (New England Biolabs, Ipswich, MA), anti-pparβ/δ (ab21209) (Abcam, San Francisco, CA), anti-pdpk1, anti-p70 S6K (BD Biosciences Pharmingen, San Diego, CA), anti-ilk (Upstate, Lake Placid, NY), RAS activity assay GTP-bound active RAS was precipitated by binding to RAF-1 Ras binding domain (RBD) conjugated agarose followed by western blot analysis using anti-hras antibody (C20) (Santa Cruz Biotechnology, Santa Cruz, CA) as recommended by manufacturer s protocol (Millipore, Billerica, MA). Briefly, cells were lysed in magnesium lysis buffer (125 mm HEPES, ph 7.5, 750 mm NaCl, 5% lgepal CA-630, 50 mm MgCl 2, 5 mm EDTA and 10% glycerol) and 1mg of cell lysates were incubated with 5 μg of Raf-1 RBD agarose for 30 minutes at 4 C. Agarose were briefly spun down and washed three times with magnesium lysis buffer 135

149 before subject to SDS-PAGE and western blot analysis with anti-hras antibody. For GTP loading assay, equal amount (500 μg) of cell lysates from HRAS-expressing wild-type and Pparβ/δ-null keratinocytes were incubated with GDP to the final concentration of 0.1 mm for 30 minutes at 30 C so that almost all the RAS was in GDP-bound form. Then GTPγS was added to the cell lysates to the final concentration of 1 mm and the reaction was incubated at 30 C for different periods of time. The reaction was stopped by putting the tubes on ice and adding MgCl 2 to the final concentration of 60 mm. GTP-bound of RAS was detected as described above Chromatin immunoprecipitation (ChIP) ChIP was performed as previously described (Palkar et al., 2010). Briefly, cells were cross-linked with formaldehyde (1%) for 10 minutes at room temperature and then glycine was added to stop the cross-linking reaction. Cells were washed with ice-cold PBS, incubated in lysis buffer for 30 minutes, and cell debris removed by centrifugation. The chromatin was sonicated to generate DNA fragments with a range of bp. Chromatin was pre-cleared with blocked protein A/G slurry (Santa Cruz Biotechnology, Santa Cruz, CA), and then equal amounts of pre-cleared chromatin was immunoprecipitated with one of the following antibodies: anti-pparβ/δ (H-74), anti-hdac1 (H-51), anti-hdac3 (H-99) (Santa Cruz Biotechnology, Santa Cruz, CA) or acetylated histone 4 (Millipore, Temecula, CA). Rabbit IgG was used as a negative control. Antibody against mouse PPARβ/δ was described previously (12). Immunoprecipitations were 136

150 washed with low salt buffer, high salt buffer and then with TE buffer before incubation with extraction buffer (0.1 M NaHCO 3, 1% SDS). NaCl was added to the final concentration of 0.2 M and crosslinking was reversed at 65 C overnight. Immunoprecipitated DNA was purified by chloroform/phenol extraction followed by ethanol precipitation and resuspension in sterile water. qpcr was performed to determine relative enrichment of specific proteins in different promoter regions and ubiquitin C genomic DNA (for mouse gene) or β-actin 3 UTR sequence (for human gene) was used for normalization. The following primers were used for ChIP analysis: Mouse Ilk PPRE1 forward primer: 5 - AAAGTTCAGGAGGTCGGATTGGGA -3 ; reverse primer: 5 - TTCTGATGGTTGTTCCCTGTCAGC -3 ; Mouse Ilk PPRE2 forward primer: 5 - TAAAGAGGCAAGCCTTCTCGCTGA -3 ; reverse primer: 5 - AAAGGGCACAGACAGTAAGCTGGA -3 ; Human Ilk PPRE1 forward primer: 5 -TGAAGACGTGGTTGTGAAGGAGAACG-3 ; reverse primer: 5 - TTTCTGTCGTGCAGGCTTCCTGT -3 ; Human Ilk PPRE2 forward primer: 5 - GCTTTGGGAGTTGCTGGCATGAAT -3 ; reverse primer: 5 - AAGCCATGATCGTCCCTTTGAGGA -3 ; Mouse Rasgpr1 promoter region forward primer: 5 -ATCCCACCGCTCTAGGTGAGAC-3 ; reverse primer: 5 -AGCCTCTCTCGCCTTGCCCA-3 ; Mouse Rasgrp1 PPRE forward primer: 5 -CCCAAAGCAAGACTAGAGGCCAAA-3 ; reverse primer: 5 -ACACCATCATTCGGAACTGGGTGA-3 ; Mouse Angptl4 PPRE forward primer: 5 - TCTGGGTCTGCCCCCACTCCTGG-3 ; reverse primer: 5 -GTGTGTGTGTGGGATACGGCTAT-3 ; Human Angplt4 PPRE forward primer: 137

151 5 - ACCGTGCAGACTCATTTCGACCTT-3 ; reverse primer: 5 - GGAGGCAGGGTTGAGGAAAGAAA-3 ; Mouse Ubc sequence forward primer: 5 -CCAGTGTTACCACCAAGAAGGTCA-3 ; reverse primer: 5 -CCATCACACCCAAGAACAAGCACA-3. Human β-actin 3 UTR sequence forward primer: 5 -AATGTGGCCGAGGACTTTGATTGC-3 ; reverse primer: 5 -AGGATGGCAAGGGACTTCCTGTAA Electrophoretic mobility shift assay (EMSA) EMSA was performed as previously described (Kim et al., 2004a). Briefly, to determine whether PPARβ/δ can directly bind to the putative PPRE identified in the murine rasgrp1 exon region, murine PPARβ/δ and RXRα were in vitro transcribed and translated (TNT, Promega) from PPARβ/δ-pcDNA and RXRα-pcDNA vectors previously described (1), and incubated with oligonucleotides encoding either the putative PPRE, or a mutated PPRE (see below), which were end labeled using [γ- 32 P] ATP in 20 μl of binding buffer containing 10 mm Tris (ph 8.0), 150 mm KCl, 0.5 mm EDTA, 0.1% Nonidet P-40, 12.5% glycerol, 0.2 mm dithiothreitol, and 0.1 μg/μl poly(di-dc). For competition, unlabeled PPRE was added at the indicated molar excess. The following oligonucleotide sequences were used: PPRE1-sense strand (5 - AAAGCAAGACTAGAGGCCAAATCAACCAAC -3 ), PPRE1-antisense strand (5 - GTTGGTTGATTTGGCCTCTAGTCTTGCTTT -3 ), mutant PPRE1-sense strand (5 - AAAGCAAGACTAGAAGCTTAATCAACCAAC -3 ), mutant PPRE1-antisense strand (5 - GTTGGTTGATTAAGCTTCTAGTCTTGCTTT -3 ), PPRE2-sense 138

152 strand (5 - CAGCTTGGCCCAGATCACCCAGTTCCGAA -3 ), PPRE2-antisense strand (5 - TTCGGAACTGGGTGATCTGGGCCAAGCTG -3 ), mutant PPRE2-sense strand (5 - CAGCTTGGCCCAGCCTTCCCAGTTCCGAA -3 ), and mutant PPRE2-antisense strand (5 - TTCGGAACTGGGAAGGCTGGGCCAAGCTG -3 ). The PPRE is underlined and the mutated bases are indicated in bold. Binding products were resolved on a 5% acrylamide gel and autoradiography was performed using standard protocols Luciferase reporter assay FOXO reporter letivirus vector (CCS-1022L) were obtained from SABiosciences (Frederick, MD). Mock infected or HRAS-expressing keratinocytes were co-infected with FOXO-luciferase and CMV-Renilla-hygromycin (CLS-RHL) control lentivirus at the estimated M.O.I of five. Puromycin and hygromycin double resistant cells were selected and luciferase was measured as previously described with a dual luciferase system (12). Rasgrp1-PPRE region was cloned with the following primers. forward primer: 5 - ATAGAGCTCGCTCCAGAGCTGGCCCC -3 ; reverse primer: 5 - ATAGCTAGCTCGGAACTGGGTGATCTGGGC -3 ; PCR product was first subcloned into TOPO vector and subsequently cloned into Sac I and Nhe I site of pgl3-tk-minipromoter vector. Mock infected or HRAS-expressing keratinocytes were transiently co-transfected with Rasgrp1-PPRE-pgl3-TK and CMV-Renilla control vector and luciferase activity was measured. 139

153 DNA microarray analysis DNA microarray analysis was essentially as previously described in chapter two. The only exception is that 72 hours of GW0742 treatment in wild-type and Pparβ/δ-null keratinocytes HRAS-expressing samples were used in this study. Gene Expression Omnibus (GEO) database ( GSE32498 is updated with raw data from these two samples Gene Set Enrichment Analysis (GSEA) GESA was performed essentially as previously described in chapter two. The following genes were included in the FOXO target gene set. Bcl2l11, Bnip3, Bcl6, Fasl, Tnfsf10, Fbxo32, G6pc, Pck1, Cdkn1A, Cdkn1b, Rbl2, Gadd45a, Ccng2, Ddb1, Sod2, Cat, Sesn1 and Btg Human adenoma samples Matched pairs of frozen normal colon tissue and colon adenomas were obtained from The Penn State Hershey Cancer Institute Tissue Bank. 140

154 3.4 Results PPARβ/δ promotes HRAS-induced senescence and suppresses malignant conversion Results from a complete carcinogen bioassay showed a higher percentage of malignant squamous cell carcinomas (SCC) and a lower percentage of benign papillomas in Pparβ/δ-null mice compared to controls (Figure 3-1A). The distribution of tumor type was not influenced by ligand activation of PPARβ/δ with GW0742 (Figure 3-1A). This suggests that PPARβ/δ may suppress malignant conversion of skin tumors. The lack of effect by an exogenous ligand could be due to the presence of high affinity endogenous ligand(s). To further characterize the role of PPARβ/δ in malignant transformation of skin tumors with an Hras mutation, an in vitro malignant conversion assay was performed. This assay uses primary keratinocytes retrovirally infected with an oncogenic HRAS vector (Roop et al., 1986) that confers a malignant phenotype characterized by resistance to calcium-induced differentiation (Morgan et al., 1992). The level of downstream pathways of HRAS, including p-erk and p-akt, are comparable to that of two skin cancer cell lines harbouring endogenous Hras mutation, therefore arguing against the supraphysilogical effects of retroviral system used in this study (Figure 3-1B). HRAS-expressing Pparβ/δ-null cells developed calcium-resistant foci (Figure 3-1C) that are bromodeoxyuridine (BrdU) positive (Figure 1D) but HRAS-expressing wild-type cells do not. Ligand activation of PPARβ/δ had no influence on foci in either genotype (Figure 3-1C). Since oncogenic HRAS could 141

155 Figure 3-1. PPARβ/δ suppresses malignant conversion. (A) Distribution of tumor types in wild-type or Pparβ/δ-null mouse skin treated with or without GW0742. (B) Western blot analysis of p-erk and p-akt in HRAS-expressing keratinocytes and 308 and SP1 cells. (C, D) After 4 weeks of culture in high Ca 2+ medium, Ca 2+ resistant foci were stained with rhodamine. The number and size of foci were quantified (C). Relative proliferating cells in the foci (BrdU positive) are shown in (D). trigger cellular senescence to prevent malignant transformation in primary cells (Serrano et al., 1997; Vijayachandra et al., 2003) and wild-type HRAS-expressing keratinocytes cultured in high calcium medium exhibited morphology reminiscent of cellular senescence (flattened cell shape and increased cytoplasm size) (Figure 3-1D), senescence-associated β-galactosidase (β-gal) and BrdU 142

156 Figure 3-2. PPARβ/δ promotes HRAS-induced senescence. (A, B) HRAS-expressing cells treated with DMSO or 1 µm GW0742 were stained for β-gal. The percentage of β-gal and BrdU positive cells was quantified. (C) β-gal staining of skin tumor samples from wild-type or Pparβ/δ-null mice. SCC: squamous cell carcinoma. SCP: squamous cell papilloma. KA: keratoacanthoma. BBCT: benign basal cell tumor. Values represent the mean ± S.E.M. * significantly different values from wild-type DMSO (P 0.05). labeling was examined. A higher percentage of BrdU labeling and lower percentage of β-gal-positive cells was noted in HRAS-expressing Pparβ/δ-null cells compared to wild-type counterparts (Figures 3-2A,B). Surprisingly, ligand activation of PPARβ/δ decreased both the percentage of β-gal- and BrdU-positive 143

157 cells in HRAS-expressing wild-type but not in Pparβ/δ-null cells (Figures 3-2A,B). 75% of papillomas from wild-type mice and 33% of papillomas from Pparβ/δ-null mice stained positive for β-gal, and no SCC from either genotype were positive for β-gal (Figures 3-2C). This is consistent with previous studies showing that benign skin papillomas undergo senescence while malignant tumors do not (Collado et al., 2005; Sun et al., 2007) PPARβ/δ promotes senescence by diverting from the PI3K/AKT to MEK-ERK signaling Downstream signaling by HRAS was examined in HRAS-expressing keratinocytes. Higher phosphorylated MEK (pmek), perk and GTP-bound HRAS was observed in HRAS-expressing wild-type cells compared to Pparβ/δ-null cells (Figures 3-3A,B). In contrast, a higher level of phosphorylated AKT (pakt) was observed in HRAS-expressing Pparβ/δ-null cells as compared to wild-type (Figures 3-3C). At a later time point, higher expression of the senescence markers p53, p21 and p27 and a decreased level of the proliferation marker phosphorylated RB (prb-s780) was observed in wild-type cells compared to Pparβ/δ-null cells (Figures 3-3D). While ligand activation of PPARβ/δ with GW0742 caused a decrease in pmek, perk, GTP-bound HRAS in HRAS-expressing wild-type cells (Figures 3-3A,B), this is likely due to the fact that ligand activated PPARβ/δ selects against cells with higher HRAS expression (Chapter two). Further, since exogenous PPAR ligands can exhibit marked differences in recruitment of co-activators as compared to an endogenous PPAR ligand (Kodera et al., 2000), comparing effects on senescence induced by 144

158 Figure 3-3. PPARβ/δ-dependent diversion from PI3K/AKT to the MEK/ERK pathway. HRAS-expressing keratinocytes were treated with DMSO or GW0742. Western blot of p-mek, MEK, p-erk, ERK (A), GTP-bound form of HRAS (B), AKT and p27 in normal or HRAS-expressing keratinocytes 5 day post HRAS expression (C) and senescence markers 11 days post HRAS expression (D). Values with different superscripts are different, P GW0742 may not be relevant in the present study. Since higher levels of pmek and perk and a lower level of pakt in wild-type cells correlates with increased senescence (Figures 3-3), the hypothesis that PPARβ/δ promotes senescence by diversion of signaling from PI3K/AKT to the MEK/ERK pathway was examined. Treating both HRAS-expressing wild-type and Pparβ/δ-null cells with the MEK1 inhibitor PD98059 delayed HRAS-induced senescence (Figures 3-4A,B). In addition, overexpression of constitutively active MEK (MEK2D) (Schramek et al., 1997) increased the percentage of SA-β-gal 145

159 Figure 3-4. p-akt suppresses and p-erk prmotes HRAS-induced senescence. Representative microphotographs (A) and percentage of SA-β-gal positive cells (B) in HRAS-expressing keratinocytes treated with different chemicals at day 2 post HRAS expression. Western blot analysis showing increased p-erk (C) and PTEN (D) level in MEK2D and PTEN overexpressing cells respectively. Representative microphotographs (E) and percentage of SA-β-gal positive cells (F) in MEK2D and PTEN overexpressing cells. # and * indicate significantly different values (P 0.05) from wild-type and Pparβ/δ-null control respectively. 146

160 Figure 3-5. PPARβ/δ-dependent diversion from PI3K/AKT to the MEK/ERK pathway. HRAS expressing wild-type and Pparβ/δ-null keratinocytes were treated with different chemicals for three days. (A) Percentage of viable cells relative to DMSO group was calculated by MTT assay (n=4). (B) Flow cytometry analysis using anti-hras antibody was performed. Pearson s second skewness coefficient, which was defined by 3x (mean-median)/standard deviation was calculated (n=3). (C) Percentage of BrdU positive cells was shown after LY treatment. # and * indicate significantly different values (P 0.05) from DMSO controls. cells in both wild-type and Pparβ/δ-null cells (Figure 3-4C,E,F). In contrast, PI3K inhibitor LY treatment or overexpression of PTEN significantly increased the percentage of SA-β-gal positive Pparβ/δ-null cells (Figure 3-4A,B, D-F). This evidence suggests that whereas higher p-mek and p-erk trigger senescence, a higher p-akt suppresses it. Consistent with the fact that Pparβ/δ-null cells are dependent on PI3K-AKT rather than MEK-ERK to evade senescence, Pparβ/δ-null cells are more sensitive to LY and less sensitive to PD98059 induced decrease of cell proliferation 147

161 (Figure 3-5A) and induced selection against higher HRAS-expressing cells (Figure 3-5B) compared to wild-type cells. In addition, LY decreased the percentage of BrdU positive HRAS-expressing wild-type and Pparβ/δ-null cells (Figure 3-5C). This evidence suggests that PPARβ/δ promotes senescence by diversion of signaling from PI3K/AKT to the MEK/ERK pathway Higher p-akt decreased FOXO activity to attenuate senescence A higher level of p27 Kip1 in normal Pparβ/δ-null cells as compared to normal wild-type cells (Figure 3-6A) argues against the previous report showing that p27 is a direct target gene that PPARβ/δ activates (Sue et al., 2009). Since p27 is positively regulated by FOXO transcription factors, whose activity is inhibited by p-akt (Greer and Brunet, 2005), FOXO activity was examined in keratinocytes. A lower FOXO activity was found in HRAS-expressing Pparβ/δ-null cells and LY treatment significant increases FOXO activity in both genotypes (Figure 3-6B,C). In addition, gene set enrichment analysis (GSEA) from a microarray dataset (Chapter two) showed lower FOXO target genes expression in HRAS-expressing Pparβ/δ-null cells (Figure 3-6D,E). LY decreased p-akt and increased senescence marker including p27 and DcR2 in HRAS-expressing keratinocytes (Figure 3-6F). This evidence suggests that decreased FOXO activity resulting from a higher AKT activity could account for the reduced senescence in Pparβ/δ-null cells. 148

162 Figure 3-6. Higher p-akt decreased FOXO activity to attenuate senescence. (A) Quantitative real-time PCR of p27 Kip1 in normal and HRAS-expressing keratinocytes treated with DMSO or 1 μm GW0742 for three days. The signal was normalized to that of β-actin (n=3) (B) Luciferase assay on FOXO transcription factors 5 day post HRAS expression (n=3). (C) HRAS expressing wild-type and Pparβ/δ-null keratinocytes were treated with DMSO or 10 μm LY and luciferase activity assay on FOXO transcription factors 11 day post HRAS expression was performed, Relative firefly luciferase activity were measured by an illuminometer and normalized to that of Renilla luciferase (n=3). (D, E) GSEA analysis on FOXO target gene expression in HRAS expressing wild-type and Pparβ/δ-null keratinocytes revealed decreased FOXO target gene expression in Pparβ/δ-null cells 5 day post HRAS expression. (F) LY decreased p-akt, increased p27, p53 and Dcr2 level in HRAS-expressing keratinocytes. Values=mean ± SEM. # indicates significantly different values (P 0.05) from wild-type DMSO control. 149

163 3.4.4 PPARβ/δ positively regulates RASGRP1 and represses ILK and PDPK1 to promote HRAS-induced senescence Microarray analysis was performed to identify PPARβ/δ target genes regulating HRAS-induced senescence. Consistent with report showing a negative feedback signaling in response to HRAS (Courtois-Cox et al., 2006), the level of negative RAS regulators RASGAP120 and RASA4 was increased and the level of positive RAS regulator RASGRP1 was decreased in response to HRAS activation (Figure 3-7A,B). Interestingly, the level of negative RAS regulators RASGAP120, RASA4, DUSP1 and DUSP 3 were higher, whereas RASGRP1 level was lower in HRAS-expressing Pparβ/δ-null cells compared to wild-type cells(figure 3-7A,B). Since RAS with mutations at 12 and 61 codon is resistant to RASGAP (Schubbert et al., 2007), it is assumed that decreased RASGRP1 expression, which is responsible for converting RAS-GDP to RAS-GTP, rather than the increased RASGAP120 level, accounts for the decreased level of RAS-GTP in Pparβ/δ-null cells. As expected, decreased GTP loading to HRAS was observed in HRAS-expressing Pparβ/δ-null cells (Figure 3-7C). This is also consistent with previous report showing that deficiency in RASGRP1 resulted in decreased level of RAS-GTP in mouse keratinocytes (Sharma et al., 2010) Examination of Rasgrp1 gene revealed two peroxisome proliferator-activated receptor response elements (PPRE) in the second exon (Figure 3-8A). ChIP analysis revealed that while HRAS led to decreased of acetylated Histone 4 (AC-H4) in both wild-type and Pparβ/δ-null cells in the promoter region, ligand activation and HRAS expression derepressed this by increasing the recruitment of 150

164 Figure 3-7. PPARβ/δ dampens HRAS-induced negative feedback response. Keratinocytes mock or HRAS-infected were treated with DMSO or GW0742 for 3d (A-H). (A) qrt-pcr of Dusp1, Rasgrp1, Rasgrp120, Rasa4 and Dusp3 (n=3). (B) Western blot of RASGAP120, DUSP1 and RASGRP1. (C) HRAS-GTP loading assays showing faster accumulation of GTP-HRAS in HRAS-expressing Pparβ/δ-null keratinocytes. Values=mean ± SEM. * indicate significantly different values (P 0.05) from HRAS-expressing wild-type DMSO control respectively. PPARβ/δ to the PPRE region, subsequently resulting in an local increase of AC-H4 in wild-type cells (Figure 3-8B). As a positive control, ligand activation and HRAS expression increased the AC-H4 level and PPARβ/δ recruitment to a well-characterized region containing multiple PPREs in Angtpl4, a PPARβ/δ target gene in wild-type cells (Mandard et al., 2004) (Figure 3-8C). None of the effects were observed in Pparβ/δ-null cells (Figure 3-8B,C). The increased binding of PPARβ/δ to Rasgrp1 PPRE, leading to an increase in enhancer activity in response to ligand and HRAS was also confirmed by luciferase and gel shift assays (Figure 3-8D,E). Integrin-linked kinase (ILK) and phosphoinositide-dependent kinase-1 (PDPK1) 151

165 Figure 3-8. Rasgrp1 is a PPARβ/δ target gene. (A, B) Diagram of Rasgrp1 promoter region and ChIP on the promoter and PPRE-containing second exon of mouse Rasgrp1 gene (n=3). (C) ChIP analysis on PPRE-containing third intron of mouse Angptl4 gene. PPREs are illustrated by three red arrows. (n=3) (D) Region containing mouse Rasgrp1 PPRE was cloned into pgl3-tk-luc vector and luciferase activity assay was performed (n=3). Both ligand activation and introduction of mutant HRAS led to an increase of luciferase activity only in wild-type cell. (E) Gel shift analysis on the two mouse Rasgrp1 PPREs. PPARβ/δ/RXRα heterodimer bind to both PPREs but they do not bind to the PPRE with mutated PPARβ/δ/RXRα dimer binding sites. * indicates significantly different values (P 0.05) from wild-type DMSO control respectively. 152

166 Figure 3-9. PPARβ/δ represses Ilk and Pdpk1. (A) qrt-pcr of Ilk and Pdpk1 in keratinocytes mock or HRAS-infected and treated with DMSO or GW0742 (n=3). (B) Western blot of ILK and PDPK1 in HRAS-expressing keratinocytes. (C) Region containing wild-type mouse Ilk PPRE (1), PPRE with mutation in the upstream PPRE half site (2) or PPRE with mutation in the downstream half site (3) were cloned into pgl3-tk-luc vector and luciferase activity assay was performed (n=3). (D, E) ChIP on the PPRE-containing second intron of mouse Ilk gene (n=3). * indicate significantly different values (P 0.05) from wild-type DMSO control respectively. are known to phosphorylate AKT at Ser473 and Thr308 respectively to fully activate AKT (Alessi et al., 1997; Persad et al., 2001) and they have been previously shown to be PPARβ/δ target genes (Di-Poi et al., 2002). However, in contrast to previous report showing that ligand activation of PPARβ/δ increases ILK and PDPK1 expression (Di-Poi et al., 2002), we found that PPARβ/δ repressed their expressions in both normal and HRAS-expressing keratinocytes 153

167 Figure PPARβ/δ overexpression decreased the mrna level of both Ilk and Pdpk1 and ligand activation of PPARβ/δ did not further increase the level of Ilk or Pdpk1 in PPARβ/δ-overexpressing HaCat cells, (A,B) Angptl4 serves as a positive control showing repression of basal ANGPTL4 mrna level by overexpression of PPARβ/δ in the absence of GW0742 and increased induction of ANGPTL4 in response to ligand activation in PPARβ/δ overexpressing cells. Western blot analysis in control migr1 (empty vector) and migr1-pparβ/δ (PPARβ/δ stably overexpressing) HaCat cell line. Overexpression of PPARβ/δ represses ILK and PDPK1 level, concomitant with decreased expression of p-akt and ER stress markers including ATF4, ATF6, IRE1α and PERK. (C) ChIP analysis on PPRE-containing second intron of human ILK gene (n=3). Consistent with mrna and protein results, in PPARβ/δ over-expressing HaCat cells, increased recruitment of PPARβ/δ is associated with increased HDAC1 and HDAC3 and decreased AC-H4 level to the upstream PPRE of Ilk gene. However, no increase in AC-H4 level or decreased HDAC1 and HDAC3 recruitment was found to the same region in response to ligand activation of PPARβ/δ in either control or PPARβ/δ-overexpressing cells. Again, Angptl4 serves as a positive control showing increased recruitment of PPARβ/δ is associated with increased recruitment of HDAC1 and HDAC3 and decreased AC-H4 to the PPRE containing region by overexpression of PPARβ/δ in the absence of GW0742. Ligand activation of PPARβ/δ in the PPARβ/δ overexpressing cells further increased the binding of PPARβ/δ and decreased the recruitment of HDAC1 and HDAC3, leading to increased AC-H4 level to the same region. * indicate significantly different values (P 0.05) from migr1 DMSO control. 154

168 Figure PPARβ/δ positively regulates RASGRP1 and represses ILK to promote senescence. Representative micrographs, quantification of percentage of SA-β-gal positive cells and western blot showing induction of senescence markers in HRAS-expressing RASGRP1 overexpressing cells (8d post HRAS) (A, B) and ILK shrna knocking-down HRAS-expressing Pparβ/δ-null keratinocytes (11d post HRAS) (C, D). ILK shrna2 serves as a negative control. * indicate significantly different values (P 0.05) from wild-type control. and ligand has no effects on their expression (Figure 3-9A,B), consistent with previous findings in our lab (Burdick et al., 2007; Marin et al., 2006). PPARβ/δ binding to the upstream PPRE in the second intron of ILK was associated with increased recruitment of HDAC1 and HDAC3 and decreased AC-H4 in the region (Figure 3-9D,E). Interestingly, while mutations in the PPARβ/δ binding half-site abolishes the repression of ILK by PPARβ/δ, mutating the downstream RXRα binding site to a consensus sequence increased the repression (Figure 3-9C), suggesting that the modest repression of ILK PPRE activity by PPARβ/δ was possibly due to a degenerative RXR α binding site. In addition, the repression of ILK by PPARβ/δ was also recapitulated in a HaCaT cell line stably overexpressing PPARβ/δ (Borland et al., 2008) (Figure 3-10). 155

169 Both overexpression of RASGRP1 and knocking-down of ILK alone in Pparβ/δ-null cells could rescue the decreased HRAS-induced senescence (Figure 3-11). This evidence also implies that both decreased MEK-ERK activity and increased p-akt activity are required to evade senescence PPARβ/δ attenuates HRAS-induced ER stress via inhibiting p-akt-mtor activity During the culture of HRAS-expressing keratinocytes, we consistently observed more vacuolized Pparβ/δ-null cells compared to wild-type counterparts (Figure 3-12A,B). Since vacuolization is a sign of endoplasmic reticulum (ER) stress, which can be induced by activating HRAS (Denoyelle et al., 2006), the extent of ER stress in HRAS-expressing cells was quantified. Increased staining of ER by ER-Tracker Blue-White DPX (Figure 3-12C,D), increased mrna (Figure 3-12E) and protein level of ER stress and unfolded protein response (UPR) markers ATF6, BiP, phospho-perk and IRE1α (Figure 3-12G) and increased spliced Xbp-1 mrna (Figure 3-12F) were observed in Pparβ/δ-null cells. The increased ER stress was not due to a difference in HRAS level (Figure 3-12G). Increased level of ER stress did not lead to increased apoptosis in HRAS-expressing Pparβ/δ-null cells as illustrated by the lack of increased expression of pro-apoptotic marker BAD and BAX (Figure 3-13A) and the lack of increased PI positive apoptotic cells (Figure 3-13B). Consistent with the enhanced ER stress in Pparβ/δ-null cells, HRAS-expressing Pparβ/δ-null cells were more sensitive to ER stress inducer tunicamycin-induced cell death (Figure 3-13C). 156

170 Figure PPARβ/δ attenuates HRAS-induced ER stress. (A, B) Representative micrographs and quantification of vacuolized HRAS-expressing keratinocytes. (C, D) Cells were stained with ER-tracker DPX and mean intensity of fluorescence signal was calculated. (E) Heat map of mrna level of twenty-three ER stress and UPR marker in both mock-infected and HRAS-infected keratinocytes. (F) Semi-quantitative PCR of spliced and unspliced form of Xbp-1 mrna. (G) Western blot of ER stress and UPR markers. * indicate significantly different values (P 0.05) from wild-type control. Next we tested the hypothesis that the exacerbated ER stress observed in HRAS-expressing Pparβ/δ-null cells was caused by a higher p-akt signaling. As expected, treating cells with LY or overexpression of PTEN 157

171 Figure PPARβ/δ attenuates HRAS-induced ER stress without affecting apoptosis. (A) Western blot of pro-apoptotic protein BAD and BAX in HRAS-expressing keratinocytes. (B) Quantification of PI positive apoptotic HRAS-expressing cells by flow cytometry analysis. (C) MTT assay of HRAS-expressing keratinocytes treated with 50ng/μl tunicamycin. The percentage of viable wild-type or Pparβ/δ-null cells was normalized to its own DMSO control. * indicate significantly different values (P 0.05) from wild-type control. Figure PPARβ/δ attenuates HRAS-induced ER stress via inhibiting p-akt activity. Representative micrographs and quantification of percentage of vacuolized cells treated with LY (A) or overexpressing PTEN (B). 158

172 Figure PPARβ/δ attenuates HRAS-induced ER stress via inhibiting p-akt-mtor activity. (A) Western blot of ER stress and UPR markers as well as mtor target genes in response to LY Representative micrographs (B) and quantification (C) of protein synthesis in HRAS-expressing cells treated with DMSO, 3nM rapamycin or 10μM LY Cells were pulsed with methionine analogue L-azidohomoalanine and newly synthesized proteins were detected by a chemoselective ligation reaction between the azide of L-azidohomoalanine and Alexa-488 alkyne. Representative micrographs (D) and quantification (E) of vacuolized cells treated with 3nM rapamycin. (F) Western blot of ER stress and UPR markers as well as mtor target genes in response to 3nM rapamycin. 159

173 significantly decreased the percentage of vacuolized cells in both genotypes (Figure 3-14). In addition, LY treatment decreased ER stress markers p-perk, ATF6, IRE1α and BiP (Figure 3-15A). p-akt can activate mammalian target of rapamycin (mtor), lead to increased protein synthesis and therefore potentially exacerbate ER stress. Consistent with this hypothesis, LY treatment decreased mtor targets phospho-p70 S6K, phosphp-s6 and phospho-4ebp1 level in both genotypes, all of which suggest decreased protein synthesis (Figure 3-15A). In addition, significantly increased protein synthesis was found in HRAS-expressing Pparβ/δ-null cells and both mtor inhibitor rapamycin and LY decreased protein synthesis in both genotypes (Figure 3-15B,C). Finally, inhibition of mtor by 3nM rapamycin attenuates ER stress in both genotypes without significantly decreasing p-akt level (Figure 3-15D-F) ER stress-associated UPR inhibits HRAS-induced senescence Contrary to report showing ER stress and UPR promotes HRAS-driven senescence in melanocytes (Denoyelle et al., 2006), SA-β-Gal positive staining correlated with decreased vacuolization in both wild-type and Pparβ/δ-null HRAS-expressing keratinocytes (Figure 3-16A,B). In addition, calcium-resistant Pparβ/δ-null cells showed massive vacuolization (Figure 3-16C). These findings suggest that ER stress and UPR prevent HRAS-induced senescence in keratinocytes. To prove this hypothesis, two main UPR pathways were selectively inhibited by shrna designed against XBP-1 or ATF4 in HRAS-expressing keratinocytes. XBP-1 and ATF4 deficiency increased the proportion of SA-β-Gal 160

174 Figure Decreased XBP-1 increased HRAS-induced cellular senescence. Primary keratinocytes were infected with Hras retrovirus as previously described. (A) Representative photomicrographs showing negative correlation between the extent of vacuolization and senescence. Cells in red box are SA-β-gal positive senescent cells and they do not show signs of vacuolization. In contrast, cells in black box are SA-β-gal negative cells and they are massively vacuolized. Quantification of cells that are either double positive for SA-β-gal and vacuolization (Vac), double negative for SA-β-gal and vacuolization, single positive for SA-β-gal or vacuolization is provided. P value is generated from Chi-square test showing statistically significant enrichment for SA-β-gal positive vacuolization negative [SA-β-gal (+), Vac (-)] and SA-β-gal negative vacuolization positive [SA-β-gal (-), Vac (+)] cells. (C) Photomicrographs of a calcium-resistant clone from HRAS-expressing Pparβ/δ-null cells. Arrow indicates a vacuolized cell. (D) Three different shrnas against murine Xbp-1 were tested in wild-type HRAS-expressing keratinocytes and arrow indicates the shrna that works the best. Representative micrographs (E), quantification of the percentage of SA-β-gal positive cells (F) and western blot (G) following Xbp-1 knocking-down. Values=mean ± SEM. * and # indicate significantly different values (P 0.05) from wild-type and Pparβ/δ-null ctrl shrna group respectively. 161

175 Figure Decreased ATF4 increased HRAS-induced cellular senescence. (A) Five different shrnas against murine Atf4 were tested in wild-type HRAS-expressing keratinocytes and arrow indicates the shrna that works the best. (B-D) Representative micrographs (B), quantification of the percentage of vacuolized and SA-β-gal positive cells (C) and western blot (D) following Atf4 knock-down. (E) Isolated calcium-resistant HRAS-expressing Pparβ/δ-null clones were infected with ctrl, Atf4, Xbp-1 or Ilk shrna encoding lentivirus and SA-β-gal positive cells were calculated five days post shrna infection. Representative micrographs were shown from clone1. Values=mean ± SEM. * and # indicate significantly different values (P 0.05) from wild-type and Pparβ/δ-null ctril shrna group respectively. 162

176 positive cells, concomitant with increased expression of senescence markers in both genotypes, although stronger effects were found in Pparβ/δ-null cells (Figure 3-16 E-G, Figure 3-17B-D). In addition, BiP (a target gene of both ATF4 and XBP-1) and p-akt levels were decreased following XBP-1 and ATF4 knockdown and no change in p-erk level was observed (Figure 3-16E-G, Figure 3-17B-D and data not shown). Furthermore, knockdown of XBP-1, ATF4 or ILK partially restored cellular senescence in 3 out 6 calcium-resistant HRAS-expressing Pparβ/δ-null monoclone cells isolated from a similar assay as shown in Figure 3-1C (Figure 3-17G). To further explore the relation between UPR and senescence, two common ER stress inducers tunicamycin and thapsigargin were used. Treating HRAS-expressing keratinocytes with increasing concentrations of both reagents lead to a dose-dependent increase of vacuolized cells and decrease of SA-β-Gal positive cells in both genotypes (Figure 3-18A,B, Figure 3-19A,B). Since more prominent effects were found in wild-type cells presumably because they have lower basal level of ER stress compared to Pparβ/δ-null cells (Figure 3-18A,B, Figure 3-19A,B), wild-type HRAS-expressing cells were used in the following studies. Tunicamycin and thapsigargin treatment increased the percentage of BrdU positive proliferating wild-type cells (Figure 3-19C), increased expression of UPR markers including BiP and IRE1α, proliferation marker PCNA and p-rb (S780) (Figure 3-18C, Figure 3-19E), and decreased senescence marker including p16 and DcR2 (Figure 3-18C, Figure 3-19E). Interestingly, both reagents also resulted in an increase of p-akt level as early as 5 day post HRAS 163

177 Figure Increased ER stress-associated UPR by thapsigargin attenuates HRAS-induced senescence. Primary keratinocytes were infected with Hras retrovirus as previously described. Representative micrographs (A), quantification of the percentage of vacuolized and SA-β-gal positive wild-type and Pparβ/δ-null HRAS-expressing cells (B) and western blot in wild-type HRAS-expressing cells (C) in response to different concentration of thapsigargin. Values=mean ± SEM. * and # indicate significantly different values (P 0.05) from wild-type and Pparβ/δ-null DMSO control respectively. expression and it persisted (Figure 3-18C, Figure 3-19D,E). They also selected against higher HRAS-expressing keratinocytes presumably through increased apoptosis (increased PARP cleavage), resulting in populations of cells with lower HRAS expression and subsequent decreased p-erk level (Figure 3-18C, Figure 3-19D). This is important to note because previous results showed that both attenuation of p-erk and potentiation of p-akt are required to evade senescence. 164

178 Figure Increased ER stress-associated UPR attenuates HRAS-induced senescence through a p-akt dependent mechanism. Representative micrographs (A) and quantification of the percentage of vacuolized and SA-β-gal positive cells (B) in response to different concentration of tunicamycin. (C) Quantification of the percentage of BrdU positive wild-type cells in response to 25 ng/μl tunicamycin, 25 ng/μl tunicamycin plus 10 μm LY or 2.5 nm thapsigargin 12 days post HRAS expression. Western blot of wild-type HRAS expressing keratinocytes treated with 25 ng/μl tunicamycin for 3 days (D) or 25 ng/μl tunicamycin or 25 ng/μl tunicamycin plus 10 μm LY for 9 days (E). (F) Quantification of the percentage of vacuolized and SA-β-gal positive cells as in (E). Values=mean ± SEM. * and # indicate significantly different values (P 0.05) from wild-type and Pparβ/δ-null DMSO control respectively. 165

179 To determine if tunicamycin attenuated senescence and increased proliferation by increasing p-akt signaling, tunicamycin was co-treated with LY to inhibit p-akt. While LY partially abolishes the effects of attenuating senescence by tunicamycin alone (Figure 3-19F), it completely abolishes the increased proliferation by tunicamycin alone (Figure 3-19C). These effects were not due to decreased ER stress (Figure 3-19E,F). These findings suggest that increased UPR attenuates senescence through both p-akt dependent and independent mechanisms. The fact that enhanced p-akt activity increase ER stress and ER stress-associated UPR in turn increases p-akt level suggests the presence of a positive feedback loop between p-akt, ER stress and UPR. If so, then a brief trigger of ER stress should be sufficient to establish the positive loop, maintain higher UPR and higher p-akt activity, leading to evasion of senescence even after the initial stimuli is removed. As expected, increased cell proliferation and decreased senescence was observed 13 days post HRAS expression when wild-type HRAS-expressing keratinocytes were treated briefly with thapsigargin for only 4 days (Figure 3-20A,B). Brief tunicamycin treatment also reduced senescence (Figure 3-20B). In addition, decreased expression of senescence marker p27, p16, p21 and DcR2, increased PCNA expression was found together with increased BiP and p-akt at later time of HRAS expression in cells briefly treated with thapsigargin (Figure 3-20C). 166

180 Figure A brief increase of ER stress-associated UPR is sufficient to attenuate HRAS-induced senescence. Quantification of the percentage of BrdU positive (B) and representative micrographs (A) and quantification (B) of the percentage of SA-β-gal positive wild-type HRAS-expressing keratinocytes (at day 13 post HRAS expression) briefly treated with 2.5 nm thapsigargin for four days. (C) Western blot of wild-type HRAS-expressing cells (day17) treated with 2.5 nm thapsigargin for four days. Values=mean ± SEM. * indicates significantly different values (P 0.05) from wild-type DMSO control. The question of how increased UPR potentiates p-akt level was addressed next. We focused on the UPR marker BiP/GRP78 for two reasons. First, expression of BiP correlated with that of p-akt whether UPR decreased or increased (Figure 3-16G, 3-17D, 3-18C, 3-19E and 3-20C). Next, in addition to the well-characterized role of ER chaperones, BiP was also found on the cell surface of cancer cells and when coupled with other cell surface protein partners it can activate PI3K-AKT pathway (Lee, 2007; Ni et al., 2011). Cell-surface BiP indeed was found in HRAS-expressing keratinocytes (Figure 3-21A) and immunoneutralizing it with an antibody (Kelber et al., 2009) lowered p-akt level 167

181 Figure UPR increases p-akt partially through cell-surface BiP. (A) Immunofluorescence in wild-type HRAS-expressing cells. Arrow indicates cell-surface BiP. (B-D) Quantification of percentage of BrdU positive (B), SA-β-gal positive cells (C) and western blot (D) of HRAS-expressing cells (day 11) treated with goat IgG or 3 μg/ml anti-bip antibody. Wild-type HRAS keratinocytes were treated with control IgG, 25 ng/μl tunicamycin plus control IgG or 25 ng/μl tunicamycin plus 3μg/ml anti-bip antbody for 11 days. Quantification of the percentage of BrdU positive (E) and SA-β-gal positive cells (F) and western blot analysis of cell proliferation and senescence markers (G). Values=mean ± SEM. * indicates significantly different values (P 0.05) from wild-type DMSO control. and significantly decreased cell proliferation concomitant with decreased PCNA and p-rb (S780) level in both wild-type and Pparβ/δ-null HRAS-expressing cells (Figure 3-21B,D). However, no change in senescence was found following immunoneutralization (Figure 3-21C). Similarly, immunoneutralizing cell-surface BiP abolished the effects of increasing cell proliferation by tunicamycin and had no obvious effects on senescence in wild-type HRAS-expressing cells. (Figure 3-21E-G). Finally, we tested the effects of increased ER stress on HRAS-expressing wild-type keratinocytes in a clonogenic assay. Introduction of oncogenic Hras into 168

182 Figure A brief increase of UPR leads to malignant conversion in vitro. (A) Clonogenic assay of wild-type HRAS-expressing keratinocytes treated with different chemicals cultured in 0.5 mm calcium medium for 4 weeks. (B) Clonogenic assay of wild-type HRAS-expressing keratinocytes treated with different chemicals cultured in low calcium medium for 4 weeks. (C) Reactive oxygen species (ROS) assay on wild-type HRAS-expressing keratinocytes treated with different chemicals for 4 days. Values=mean ± SEM. * indicates significantly different values (P 0.05) from DMSO control. primary keratinocytes led to a complete senescence response and virtually no colony was found after 4 weeks of culture in either low calcium or 0.5 mm calcium medium (Figure 3-22A,B). Decreasing p-erk activity by PD98059 alone had no effects (Figure 3-22A,B). However, briefly treating cells with 1 nm or 2.5 nm thapsigargin alone resulted in a considerable number of colonies in both low calcium and 0.5 mm calcium medium (Figure 3-22A,B). Interestingly, continued treatment of 2.5 nm thapsigargin resulted in no colonies cultured in low calcium and far fewer number of colonies in 0.5 mm calcium medium after 4 weeks, even 169

183 though a number of colonies were initially observed at early times of thapsigargin treatment but they eventually sloughed off (Figure 3-22A,B and observation). Increased ER stress may cause an increase of reactive oxygen species (ROS) level (Hsieh et al., 2007), potentially increasing the chance of secondary cooperating mutations, which may lead to evasion of senescence. To rule out this possibility, the level of ROS was measured in wild-type HRAS-expressing keratinocytes after four days of thapsigargin treatment. No change in ROS level was found (Figure 3-22C), therefore it is unlikely that cooperating mutations contributes to evasion of senescence. In addition, enhanced UPR was also essential for evasion of senescence in Pparβ/δ-null HRAS-expressing cells as immunoneutralizing cell-surface BiP with anti-bip antibody completely abolished the clonal expansion of calcium-resistant foci in a similar clonogenic assays (data not shown) PPARβ/δ attenuates p-akt and ER stress to promote senescence in mouse skin tumors in vivo To test our hypothesis in vivo, chemically-induced skin tumors from one stage bioassay were examined first. Since ligand activation of PPARβ/δ has little effects on senescence, GW0742 treated mice were not examined. While SA-β-Gal positive regions of senescence papillomas are stained positive for p27 and negative for p-akt (S473) in both wild-type and Pparβ/δ-null mice (Figure 3-23A), squamous cell carcinomas exhibited strong expression of p-akt (S473) and essentially no p27 (Figure 3-23B) expression. These findings are consistent with 170

184 Figure PPARβ/δ promotes cellular senescence in mouse skin tumors in vivo. Immunofluorescene of p-akt (S473), KERATIN 14 and p27 in benign papillomas (A) and squamous cell carcinomas (B). p27 positive regions were also stained positive for SA-β-gal. (C) Immunofluorescence of p-akt (S473) and Ki-67 in skin tumors. 171

185 Figure PPARβ/δ attenuates ER stress in mouse skin tumors in vivo. (A) Representative immunofluorescence micrographs and scatter plot of p-akt (S473) and ATF4 mean intensity. Correlation coefficient (r) and p value showing how significant the slope is bigger than zero are shown. (B) Cumulative frequency of mean intensity of p-akt (S473) and ATF4 from at least 1000 cells. the role of p-akt-foxo-p27 in HRAS-induced senescence in primary keratinocytes. Ki-67 is a marker of cell proliferation and Ki-67 positive lesions were found in 4 out of 4 tumors from Pparβ/δ-null mice and essentially absent in 3 of 3 tumors examined from wild-type mice (Figure 3-23C), The fact that Ki-67 staining is absent even in non-senescent wild-type tumors suggest that PPARβ/δ substantially restrained the proliferative capacity of wild-type skin tumors. Next we examined the relation between ER stress and p-akt in vivo. p-akt (S473) level positively correlated with ATF4 and XBP-1 expression in both 172

186 Figure ER stress and UPR negatively correlates with cellular senescence in mouse skin tumors in vivo. (A) Representative immunofluorescence micrographs of p-akt (S473), XBP-1 and DcR2 in normal mouse skin and tumors. (B) Increased nuclear to cytoplasm ratio of XBP-1 in skin tumors compared to normal skin. (C) Scatter plot of p-akt (S473) and XBP-1 mean intensity in skin tumors from at least 1000 cells. Correlation coefficient (r) and p value showing how significant the slope is bigger than zero are shown. Representative immunofluorescence micrographs of p-akt (S473), XBP-1 and DcR2 in papillomas from wild-type (D) and Pparβ/δ-null (E) mice. Magnified regions enclosed in different color were shown with the original region of the same color. White arrows indicate cells with lower p-akt, higher DcR2 and higher nuclear XBP-1 level and yellows arrows indicates cells of the opposite. (F) Quantification of nuclear to cytosol ratio of XBP-1 in both senescent (DcR2 positive and p-akt negative) and proliferating regions (DcR2 negative and p-akt positive region). 173

187 wild-type and Pparβ/δ-null mice skin tumors (Figure 3-24A,3-25C). In addition, higher p-akt (S473) and higher nuclear ATF4 level was found in skin tumors from Pparβ/δ-null mice (Figure 3-24B), suggesting Pparβ/δ-null mice skin tumors exhibited a higher level of ER stress. XBP-1 and DcR2 were chosen as ER stress and senescence markers to examine the relation between ER stress and senescence in vivo. During increased ER stress, xbp-1 mrna is alternatively spliced, resulting in nuclear translocation of active XBP-1, therefore an increased ratio of nuclear to cytoplasm XBP-1 indicates increased ER stress (Carrasco et al., 2007; Hsieh et al., 2007). DcR2 is a senescence marker highly expressed in senescent skin papillomas but not in normal skin (Collado et al., 2005). Increased nuclear to cytoplasm ratio of XBP-1 was found in skin tumors compare to normal skin in both wild-type and Pparβ/δ-null mice and significantly higher nuclear to cytoplasm ratio was found in Pparβ/δ-null tumors compared to wild-type ones (Figure 3-25A,B), consistent with increased ER stress observed in Pparβ/δ-null tumors shown by ATF4 staining (Figure 3-24B). In addition, stronger p-akt (S473) staining correlated with increased nuclear XBP-1 staining in non-senescent regions. In contrast, senescent regions positively stained for DcR2 (and also positive for SA-β-Gal) are essentially negative for p-akt (S473) and more importantly showed increased cytoplasm XBP-1 distribution (Figure 3-25D-F). These effects were observed in both wild-type and Pparβ/δ-null mice skin tumors even though more striking correlation was observed in Pparβ/δ-null mice (Figure 3-25D-F). Similar correlations between p-akt, ER stress and senescence were also 174

188 Figure PPARβ/δ attenuates p-akt and ER stress to promote senescence in vivo. Western blot analysis and linear regression of p-akt and different senescence and ER stress markers in mouse skin tumor samples. Samples from Pparβ/δ-null mice are shown in red. The level of different proteins is normalized to that of LDH. confirmed by quantitative western blot analysis from both wild-type and Pparβ/δ-null mice skin tumors (Figure 3-26). In addition, higher level of ILK was also found in skin tumors from Pparβ/δ-null mice (Figure 3-26). These findings highly suggest the anti-tumorigenic role of PPARβ/δ in promoting senescence and the new pro-tumorigenic function of ER stress in attenuating senescence in vivo PPARβ/δ promotes and UPR attenuates cellular senescence in human benign lesions. Next, we focused upon finding human in vivo correlates of our observations made in mouse models. First, we analyzed the correlation between PPARβ/δ and senescence marker p16 in human benign dermal neurofibroma lesions. We choose this lesion for two reasons. First, benign dermal neurofibroma harbors Nf1 loss-of-function mutations, resulting in activation of RAS signaling pathway 175

189 Figure PPARβ/δ promotes and UPR attenuates cellular senescence in human benign lesions. (A) mrna level of p16 in different neurofibroma tumor types and tumor-derived cell lines. (B) Plot of log2 value of PPARβ/δ mrna with that of p16, RASGRP1 and PDPK1 in NF1-derived primary benign neurofibroma 176

190 Schwann cells with Nf1-/- mutation. (C) Western blot analysis of paired normal human colon and human colon adenoma. Protein level was normalized to that of ponceau staining intensity. (D) SA-β-Gal staining and immunofluorescence of p16 and BiP in human adenomas serial sections. (E) Negative correlation between p53 and three ER stress markers in human colon ademona samples. Note that p53 level is increased in adenomas as compared to normal colon, suggesting p53 is a robust senescence marker. Correlation coefficient (r) and p value showing how significant the slope is smaller than zero are shown. (Courtois-Cox et al., 2006), similar to our activated HRAS model. Secondly, SA-β-Gal positive staining senescence cells were found in these lesions (Courtois-Cox et al., 2006). Due to the unavailability of human tissue samples, we analyzed one public available microarray database related to neurofibroma (Miller et al., 2009). First, we tested the robustness of using p16 as the senescence marker. p16 mrna level is significantly higher in benign dermal neurofibroma and NF1-derived primary benign neurofibroma Schwann cells compared to malignant peripheral nerve sheath tumour (MPNST) and MPNST cell lines respectively (Figure 3-27A), implying p16 is a robust senescence marker in these type of lesions. Since neurofibromas is a hetereogenous tumor that consists not only of Schwann cells with initiating Nf1-/- mutations, but also recruited fibroblasts, peripheral cells, neurons and mast cells, of which a large percentage are merely heterozygous for an NF1 mutation and the fact that SA-β-Gal positive staining were only found in cells with homozygous loss-of-function Nf1 mutation (Courtois-Cox et al., 2006), we focused on eight samples of NF1-derived primary benign neurofibroma Schwann cells with Nf1-/- mutation. As expected, strong positive correlation between PPARβ/δ and p16 was found in these cells (Figure 3-27B). In addition, positive correlation was also found between PPARβ/δ and RASGRP1, a correlation supported by our mouse model (Figure 3-27B). While no 177

191 correlation between PPARβ/δ and ILK was found, a negative correlation between PPARβ/δ and PDPK1 was found in these cells (Figure 3-27B). Next, we turned our attention to human colon adenomas. This lesion was chosen because it contains cells displaying both strong p16 immunopositivity and absence of both mitotic and Ki-67 positivity as previously reported (Dai et al., 2000; Kuilman et al., 2008). Of the five human colon adenomas that harbor Kras mutation at codon 13, PPARβ/δ protein level was higher in colon adenomas compared to paired normal colon (Figure 3-27C). In addition, PPARβ/δ protein level in both normal colon and colon adenomas negatively correlates with both mrna and protein level of ILK and p-akt (Figure 3-27C). PPARβ/δ protein level in colon adenomas also positively correlates with that of senescence markers (Figure 3-27C). Moreover, 3 out of 5 tumors examined showed strong p16 immunopositivity and SA-β-Gal positive staining in patches of the lesions and negative correlation between p16 and BiP immunopositivity was found in all three tumors (Figure 3-27D). Negative correlation between senescence marker p53 and three UPR markers ATF6, BiP and ATF4 was also found in a human colon adenoma microarray dataset (Figure 3-27E) (Sabates-Bellver et al., 2007). This evidence strongly suggests that PPARβ/δ promotes and UPR attenuates cellular senescence in human benign lesions. 178

192 3.5 Discussion PPARβ/δ promotes HRAS-induced senescence by potentiating p-erk and attenuating p-akt activity In this study, we showed that Pparβ/δ-null keratinocytes evade HRAS-induced senescence by switching from RAF/MEK/ERK to PI3K-AKT pathway in HRAS-induced neoplastic transformation. More importantly, we demonstrated that both attenuation of p-erk and increase of p-akt activity are required to evade cellular senescence. Several studies have reported that higher p-erk activity can trigger senescence by upregulating expression of p16 Ink4a, p21 cip1, p19 ARF and p53 (de Keizer et al., 2010; Palmero et al., 1998; Satyanarayana et al., 2004; Zhu et al., 1998). On the other hand, PI3K-AKT pathway has been shown to prevent HRAS-induced senescence by preventing HRAS-induced autophagy and inhibiting FOXO transcription factor activity and increasing MDM2 activity, therefore indirectly reducing expression of p27, p21 and p53 (Courtois-Cox et al., 2006; Kennedy et al., 2011), all of which are known senescence inducers (de Keizer et al., 2010; Lin et al., 2010; Serrano et al., 1997). Decreased p27 and p21 expression was most likely due to decreased FOXO activity observed in HRAS-expressing Pparβ/δ-null keratinocytes. In addition, lower p53 expression could be well explained by the higher phospho-mdm2 level observed in Pparβ/δ-null cells (data not shown). PI3K-AKT signaling can also counteract cellular senescence through stimulating cell proliferation by phosphorylating a plethora of substrates (reviewed in (Manning and Cantley, 2007)). However, the 179

193 fact that inhibition of p-akt significantly decreased cell proliferation while had little effects on senescence in wild-type HRAS-expressing cells implied that there might be different threshold levels of p-akt required for exerting pro-proliferation and anti-senescence functions PPARβ/δ dampens the negative feedback response to potentiate p-erk signaling. It has been shown recently that mutations affecting NF1, Raf, and Ras induced a global negative feedback response that potently suppresses Ras to promote senescence by subsequent inhibition of p-akt activity (Courtois-Cox et al., 2006), a phenomenon also observed in present studies. Interestingly, PPARβ/δ dampens this negative response without increasing p-akt levels. Therefore, it is proposed that Pparβ/δ-null keratinocytes evades Hras-induced cellular senescence utilizing a strategy that involves (1) strengthening the negative feedback response to further attenuate p-erk activity and (2) compensating for the possible decreased p-akt activity by increasing ILK and PDPK1 expression. We demonstrated that PPARβ/δ de-repressed Rasgrp1 expression by directly binding to PPREs and it would be interesting to see if PPARβ/δ de-activated Rasgap120, Dusp1 and Dusp3 expression in response to Hras through a similar PPRE-dependent mechanism. 180

194 3.5.3 PPARβ/δ represses ILK and PDPK1 expression One mechanism proposed to explain the pro-tumorigenic effect of PPARβ/δ suggested that ligand activation of PPARβ/δ increases the expression of PDPK1 and ILK, causing increased phosphorylation of AKT, leading to anti-apoptotic signalling and enhanced cell survival (Di-Poi et al., 2002). However, results from our and other labs showed expression of PDPK1 and ILK is not increased and/or phosphorylation of AKT is not increased by ligand activation of PPARβ/δ, despite upregulation of known PPARβ/δ target genes (Borland et al., 2008; Burdick et al., 2007; Narkar et al., 2008; Szeles et al., 2010; Tachibana et al., 2005). In contrast, PPARβ/δ repressed ILK and PDPK1 expression, a finding also consistent with previous studies (Burdick et al., 2007; Marin et al., 2006; Yang et al., 2010). PPARβ/δ is known to repress target genes through physical interaction with co-repressors such as HDAC and SMRT (Shi et al., 2002). Repression of ILK by PPARβ/δ is associated with direct binding of PPARβ/δ and increased recruitment of HDAC1 and HDAC3 to its PPRE. So far we have not been able to identify functional PPRE within ± 5 kb region of either mouse nor human pdpk1 promoter, therefore it is possible that PPARβ/δ represses Pdpk1 through a PPRE-independent mechanism Effects of ligand activation of PPARβ/δ on HRAS-induced senescence Ligand activation of PPARβ/δ by an exogenous ligand GW0742 delayed HRAS-induced senescence. The counter-intuitive effects of exogenous ligand on senescence were due largely to the selection against higher HRAS-expressing 181

195 exerted by the ligand (Zhu et al., 2012). While ligand activation of PPARβ/δ does not change pakt signaling, ligand activated PPARβ/δ can both increase and decrease perk signaling. Ligand activation of PPARβ/δ can increase perk signaling through transcriptional upregulation of RASGRP1 in HRAS-expressing cells. On the other hand, it also decreases perk level through selection against higher HRAS-expressing cells. In the absence of exogenous ligand as found in control cells, the effects of increasing perk was greater than that of decreasing perk by potential endogenous ligand, causing a net increase of perk signaling as compared to Pparβ/δ-null cells. However, ligand activation of PPARβ/δ by GW0742 caused a much greater decrease of perk signaling due to the selection, which offsets the effects of increased RASGRP1 expression on perk signaling. As a result, the outcome is a decrease of perk signaling and delayed senescence. The differential effects between endogenous and exogenous ligand may reflect different levels of ligand needed (lower amount of ligand needed to upregulate Rasgrp1 versus higher amount of ligand needed to exert selection against higher Hras-expressing cells) regulating these two different events (direct transcriptional regulation of RASGRP versus cross talk with p107/p130). In addition, it may also be caused by marked differences in recruitment of co-activators for these two ligands (Kodera et al., 2000). Further, the possibility that the increased expression of Rasgrp1 by GW0742 in HRAS-expressing cells was in part due to the dampened negative feedback loop caused by the decreased expression of HRAS cannot be ruled out. Actually, decreased mrna level of Rasgap120 and Rasa4 in response to GW0742 in wild-type 182

196 HRAS-expressing cells could also be caused by it. However, it is worth pointing out that PPARβ/δ does transcriptionally regulate Rasgrp1, since ligand activation of PPARβ/δ increased recruitment of PPARβ/δ to the PPRE and increased luciferase activity in a reporter assay even in normal wild-type cells ER stress-associated UPR attenuates HRAS-induced senescence Increased ER stress have been observed in a variety of tumor types and ER stress associated UPR has been known to play pro-survival roles that include preventing apoptosis, promoting cell proliferation, metastasis, and allowing dormant cancer cells to survive drug therapy (reviewed in (Lee and Hendershot, 2006)). Here, we demonstrated a new pro-carcinogenic function of UPR on attenuating HRAS-induced senescence. Increased ER stress selects against higher HRAS-expressing cells, leading to decreased p-erk activity. In addition, heightened UPR increased phosphorylation of AKT partially through a cell-surface BiP/GRP78 dependent mechanism. Therefore, both of the criterions for evasion of senescence have been met. Increased p-akt would also exacerbate ER stress through increased mtor activity, completing a positive feedback loop between ER stress, UPR and.p-akt. Meanwhile, negative feedback from UPR can also hold ER stress in check, preventing unlimited amplification of the positive feedback loop. More importantly, a brief increase of ER stress by tunicamycin or thapsigargin is sufficient to establish the positive loop and lead to evasion of senescence in a subpopulations of cells in vitro and malignant conversion in vivo. We propose that only cells with the perfect balance of every component 183

197 mentioned above can evade senescence. Loss of PPARβ/δ gives HRAS-expressing cells an advantage of evading senescence by lowering p-erk activity, increasing p-akt and ER stress-associated UPR (Figure 3-28). Our model also suggests that the tumor-promoting property of thapigargin may depend on the senescence attenuating effect of UPR (Lowry et al., 1996). UPR increases the phosphorylation of AKT through a cell-surface BiP dependent mechanism. Interestingly, inhibition of cell-surface BiP by immunoneutralization decreased cell proliferation while having no effects on cellular senescence. One possible explanation is that the decrease of p-akt by immunoneutralization is relatively subtle compared to the one by LY and cell proliferation is more sensitive to the change of p-akt level than senescence. It is also likely that immunoneutralizing cell-surface BiP triggers p-akt independent responses that offset the pro-senescence effects of decreased p-akt signaling. BiP is unlikely to be the sole effecter of increasing p-akt signaling. XBP-1 was also shown to be able to increase p-akt level through an IGF dependent mechanism (Hu et al., 2007). In addition, increased expression of UPR markers PERK in response to increased ER stress also can increase PI3K/AKT signaling (Ekaterina Bobrovnikova-Marjon et al., 2012; Hu et al., 2007). Increased ER stress may also attenuate senescence independent of p-akt. Recently, an elegant study showed that ATF4 promoted tumorigenesis by directly repressing p16 ARF and p19 expression (Horiguchi et al., 2011). Therefore, ER stress-associated UPR can facilitate cellular senescence evasion through multiple mechanisms (Figure 3-28). 184

198 Figure A model of regulation of Hras-induced senescence by PPARβ/δ Anti-tumorigenic role of PPARβ/δ in human cancer The role of PPARβ/δ in human tumorigenesis remains controversial because of disparities in literature. (reviewed in (Peters et al., 2012)). Here we demonstrated that PPARβ/δ mrna level positively correlates with that of p16 and RASGRP1 in human benign dermal neurofibromas. These findings are consistent with results obtained from our mouse models with activated Hras mutation. Due to limited samples size in this microarray database, a more thorough investigation of the relation between PPARβ/δ and p16, especially at the protein level in human dermal neurofibroma tissues is necessary. In colon samples, we found that PPARβ/δ expression is increased in benign adenomas compared to normal tissue, a finding consistent with previous results (Yang et al., 2010; Yang et al., 2011a) and higher PPARβ/δ expression correlates with higher level of senescence markers in these lesions. In addition, previous reports showed that PPARβ/δ expression is decreased in malignant adenocarcinomas (Foreman et al., 2011; 185

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206 4.1 Abstract Ligand activation of peroxisome proliferator-activated receptor-β/δ (PPARβ/δ) and inhibition of cyclooxygenase-2 (COX2) activity by non-steroidal anti-inflammatory drugs (NSAID) can both attenuate skin tumorigenesis. The present study examined the hypothesis that combining ligand activation of PPARβ/δ with inhibition of COX2 activity will increase the efficacy of chemoprevention of chemically-induced skin tumorigenesis over that observed with either approach alone. To test this hypothesis, wild-type and Pparβ/δ-null mice were initiated with 7, 12-dimethylbenz[a]anthracene (DMBA), topically treated with 12-O-tetradecanoylphorbol-13-acetate (TPA) to promote tumorigenesis, and then immediately treated with topical application of the PPARβ/δ ligand GW0742, dietary administration of the COX2 inhibitor nimesulide, or both GW0742 and nimesulide. Ligand activation of PPARβ/δ with GW0742 caused a PPARβ/δ-dependent delay in the onset of tumor formation. Nimesulide also delayed the onset of tumor formation and caused inhibition of tumor multiplicity (46%) in wild-type mice but not in Pparβ/δ-null mice. Combining ligand activation of PPARβ/δ with dietary nimesulide resulted in a further decrease of tumor multiplicity (58%) in wild-type mice but not in Pparβ/δ-null mice. Biochemical and molecular analysis of skin and tumor samples demonstrate that these effects were due to modulation of terminal differentiation, attenuation of inflammatory signaling and induction of apoptosis, through both PPARβ/δ-dependent and PPARβ/δ-independent mechanisms. Increased levels and activity of PPARβ/δ by nimesulide was also observed. These studies support the hypothesis that 193

207 combining ligand activation of PPARβ/δ with inhibition of COX2 activity increases the efficacy of preventing chemically-induced skin tumorigenesis as compared to either approach alone. 4.2 Introduction Cyclooxygenase (COX) signaling pathways have important roles in modulating skin carcinogenesis. COX is the central enzyme in prostanoid biosynthesis that catalyzes the conversion of arachidonic acid to prostaglandin H 2, which is then converted to biologically active lipids such as thromboxane (TXA 2 ), prostaglandin E 2 (PGE 2 ) and prostacyclin (PGI 2 ) by different enzymes (Smith, 1989). There are two isoforms of COX, COX1 and COX2. While COX1 is constitutively expressed, COX2 is induced by tumor promoters, growth factors and cytokines (Hla et al., 1993). Results from experimental animal models have established a causal relationship between COX2 and skin carcinogenesis. For example, genetic disruption of both COX1 and COX2 can prevent skin tumorigenesis (Tiano et al., 2002) and non-steroidal anti-inflammatory drugs (NSAIDS) that inhibit COX activity inhibit both UV-induced and chemically-induced skin carcinogenesis (Bility et al., 2010; Fischer et al., 2003; Kim et al., 2006b; Pentland et al., 1999). The proliferative effects of COX2 are due primarily to increased synthesis of prostaglandins (PGs), which directly influence cell growth after binding to specific cell surface receptors, including the prostaglandin E (EP), prostaglandin F (FP) and prostaglandin I (IP) class of receptors (Hata and Breyer, 2004; Tober et al., 2007). For example, pro-tumorigenic effect of PGE 2 can be mediated by the EP2 194

208 receptor (Sung et al., 2005). While PGs can mediate their biological effects through specific prostaglandin receptors like EP, FP and IP, PGs might also modulate the activities of peroxisome proliferator-activated receptors (PPAR). Three distinct isoforms, PPARα, PPARβ (also referred to as PPARδ or PPARβ/δ) and PPARγ exist with essential roles in the regulation of adipogenesis, lipid metabolism, cell proliferation/apoptosis, cell differentiation, inflammatory responses and carcinogenesis (Burdick et al., 2006; Lalloyer and Staels, 2010; Lee et al., 2003; Peraza et al., 2006; Peters and Gonzalez, 2009; Peters et al., 2008). PPARs regulate these pathways by modulation of gene expression through direct and indirect mechanisms. PPARβ/δ is found at very high levels in the nucleus of epithelium including intestine and in keratinocytes (Girroir et al., 2008b). In the absence of ligands, nuclear PPARβ/δ can also be co-immunoprecipitated with its heterodimerization partner RXRα, suggesting that PPARβ/δ has an important constitutive role in the epithelium (Girroir et al., 2008b). Thus, it is not surprising that important roles for PPARβ/δ have been observed in skin. For example, Pparβ/δ-null mice exhibit enhanced epidermal hyperplasia in response to phorbol ester treatment (Michalik et al., 2001; Peters et al., 2000) and exacerbated chemically-induced skin tumorigenesis in a two stage carcinogen bioassay as compared to wild-type mice (Kim et al., 2004b), suggesting that PPARβ/δ inhibits epidermal cell proliferation in response to stimuli. Consistent with this idea, PPARβ/δ-dependent inhibition of skin tumorigenesis is found after topical application of the PPARβ/δ ligand GW0742 (Bility et al., 2008). The chemopreventive effects of ligand activation of PPARβ/δ are mediated in part by 195

209 induction of unidentified target genes or non-transcriptional events that modulate terminal differentiation and inhibit cell proliferation and/or inhibition of pro-inflammatory signaling (reviewed in (Burdick et al., 2006; Peters and Gonzalez, 2009; Peters et al., 2008)). Some reports suggest that NSAIDs attenuates carcinogenesis by inhibiting PPARβ/δ expression and/or activities although this view has yet to be experimentally confirmed and there are many inconsistencies with this hypothesis in the literature (reviewed in (Peters and Gonzalez, 2009; Peters et al., 2008)). For example, the hypothesis that NSAIDs inhibit cancer by decreasing PPARβ/δ expression/function is inconsistent with the observation that PPARβ/δ expression following exposure to NSAIDs is either unchanged or increased in human cancer cell lines (Foreman et al., 2009). Further, inhibition of chemically induced skin tumorigenesis is found in both wild-type and Pparβ/δ-null mice following treatment with the COX1/COX2 inhibitor sulindac, suggesting that NSAIDs mediate chemoprevention of chemically-induced skin tumorigenesis through PPARβ/δ-independent mechanisms (Kim et al., 2006b). This is consistent with a recent report showing that combining COX2 inhibition with ligand activation of PPARβ/δ resulted in increased efficacy in the inhibition of pre-existing skin tumor multiplicity (Bility et al., 2010). Collectively, these observations suggest that combining these two therapeutic approaches will increase the efficacy of chemoprevention as compared to either agent alone. Thus, the effect of combining COX2 inhibition and ligand activation of PPARβ/δ on chemoprevention of skin carcinogenesis was examined. 196

210 4.3 Materials and Methods Two-stage chemical carcinogenesis bioassay Female wild-type and Pparβ/δ-null mice on a C57BL/6 genetic background (Peters et al., 2000b), 6~8 weeks of age, were initiated with 50 μg of 7,12-dimethylbenz[a]anthracene (DMBA; Sigma-Aldrich, St Louis, MO). One week after initiation, mice were treated topically with 5 μg of 12-O-tetradecanoylphorbol-13-acetate (TPA; NCI Chemical Carcinogen Reference Standard Repository), 3 days/week for forty-one weeks. Mice from both genotypes were randomly divided into one of the following four groups: 1) control diet and topical application of acetone, 2) control diet and topical application of GW0742 (5 μm), 3) nimesulide diet (400 mg/kg) and topical application of acetone, or 4) nimesulide diet (400 mg/kg) and topical application of GW0742 (5 μm). Since C57BL/6 mice weighing grams typically consume approximately 4 grams of food per day (Bachmanov et al., 2002), the estimated dose of nimesulide ranged from mg/kg body weight per day. The concentrations of topical GW0742 and nimesulide in the diet were based on previous work showing inhibition of chemically-induced skin tumorigenesis by GW0742 or nimesulide in related models (Bility et al., 2008; Bility et al., 2010). After forty-two weeks, mice were euthanized by overexposure to carbon dioxide. Tumor samples were either fixed or snap frozen in liquid nitrogen for future analysis. Fixed tumor samples were embedded in paraffin, sectioned and stained 197

211 with hematoxylin and eosin (HE) and scored for benign or malignant pathology by two independent pathologists Short-term bioassay Female wild-type and Pparβ/δ-null mice were acclimated to either a control or nimesulide diet (400 mg/kg) for one week and then treated topically with acetone or TPA dissolved in acetone (5 μg) followed one hour later by topical application of either acetone or GW0742 (5 μm) every other day for a total of three applications. Mice were fed either the control or nimesulide diet during this period of topical GW0742 treatment. Mice were euthanized 6 hours after the last acetone or GW0742 treatment and skin samples were obtained for RNA and protein isolation Keratinocyte culture Primary mouse keratinocytes were isolated from 2-day postnatal wild-type and Pparβ/δ-null mice as described previously (Dlugosz et al., 1995). Keratinocytes were cultured in low calcium (0.05 mm) Eagle s minimal essential medium with 8% chelexed fetal bovine serum at 37 C and 5% carbon dioxide Caspase 3/7 activity assay Skin samples were ground to a fine powder in liquid nitrogen and then homogenized in buffer containing 10 mm Tris (ph 7.5), 100 mm NaCl, 1 mm 198

212 EDTA, 0.01% Triton-X100. For in vitro analysis of caspase 3/7 activity, primary keratinocytes were cultured as described above for two days before treatment with either DMSO, 1 μm GW0742, 500 μm nimesulide, or the combination of 1 μm GW0742 and 500 μm nimesulide for 24 hours. Cells were then trypsinized and lysed in the Tris buffer described above for 30 min on ice. Homogenates were centrifuged at 16,000 g, and the supernatant was used for analysis. Caspase 3/7 activity was measured using a luminescent assay (Promega, Madison, WI) Western blot analysis Primary keratinocytes were cultured as described above for two days before treatment with either DMSO, 1 μm GW0742, 500 μm nimesulide or the combination of 1 μm GW0742 and 500 μm nimesulide for 24 hours. Cells were then trypsinized and then lysed in buffer containing protease inhibitors. Samples were sonicated to facilitate cell lysis before centrifugation at 16,000 g at 4 C for 30 min and the supernatant was used for western blot analysis. Protein from skin samples was isolated similarly with the same buffer. Separation of proteins by electrophoresis, transfer to membranes and blocking was performed as previously described (Palkar et al., 2010). After incubation overnight at 4 C with the primary antibody, membranes were incubated with biotinylated secondary antibody (Jackson ImmunoResearch Laboratories, West Grove, PA) for one hour at room temperature followed by incubation with 125 I-labeled streptavidin. Membranes were exposed to plates and the level of radioactivity quantified with 199

213 filmless autoradiographic analysis. Hybridization signals for specific proteins were normalized to the signal for the loading control lactate dehydrogenase (LDH) or ACTIN. The following primary antibodies were used: anti-parp (Cell Signaling Technology, Danvers, MA), anti-k1 (Covance, Berkeley, CA), anti-k10 (Covance, Berkeley, CA), anti-pparβ/δ (Girroir et al., 2008b), anti-actin (Rockland, Gilbertsville, PA) and anti-ldh (Rockland, Gilbertsville, PA). The ratio of cleaved PARP to uncleaved PARP was calculated using Optiquant software RNA isolation and quantitative real-time PCR (qpcr) analysis Total RNA was isolated from skin and tumor samples using TRIZOL reagent (Invitrogen, Carlsbad, CA). Reverse transcription and qpcr was performed as previously described (Palkar et al., 2010). Primers for keratin 1 (K1), keratin 10 (K10), angiopoetin-like protein 4 (Angptl4), interleukin 6 (Il6) and tumor necrosis factor-α (Tnfα) have been previously described (Bility et al., 2008; Bility et al., 2010; Murray et al., 2010; Shan et al., 2008). The relative level of mrna was normalized to that of glyceraldehyde 3-phosphate dehydrogenase (Gapdh) or 18s RNA levels Statistical analysis The significance of tumor incidence between each treatment and genotype was determined by Chi-square test for trend analysis (Prism 5.0, GraphPad Software, 200

214 Inc., La Jolla, CA). Fisher s exact test was used to determine the significance of the incidence of mice with keratoacanthomas and/or squamous cell carcinomas (SCC). For all other analysis, a one-tailed student t-test was used. 4.4 Results Ligand activation of PPARβ/δ and inhibition of COX2 enhances chemoprevention of chemically-induced skin tumorigenesis Combining ligand activation of PPARβ/δ with COX2 inhibition results in a modest decrease of multiplicity of pre-existing tumors in a chemotherapeutic model (Bility et al., 2010). Since later stage tumors can be resistant to therapies designed to regress tumor growth, the effect of combining ligand activation of PPARβ/δ with COX2 inhibition was examined in a chemoprevention model. Marked changes were observed in both genotypes (Figure 4-1A). The onset of papilloma formation was sooner and the incidence of papilloma was greater in control Pparβ/δ-null mice compared to control wild-type mice prior to week 16 of the two-stage bioassay (P 0.05; Figure 4-1A), consistent with previous studies (Bility et al., 2008; Bility et al., 2010; Kim et al., 2004). Topical application of the PPARβ/δ ligand GW0742, or dietary nimesulide, caused a delay in the onset of tumor formation (P 0.05; Figure 4-1A). These effects were not found in Pparβ/δ-null 201

215 Figure 4-1. Chemoprevention of chemically-induced skin tumorigenesis by combining ligand activation of PPARβ/δ and inhibition of COX2. Wild-type (+/+) and Pparβ/δ-null ( / ) mice were treated with topical GW0742 (5 µm), dietary nimesulide (400 mg/kg) or the combination of GW0742 and nimesulide during a forty-two week two-stage bioassay (initiation with DMBA and promotion with TPA) as described in Methods. A, The incidence and onset of skin tumor formation. B, Skin tumor multiplicity. C, The average tumor size per mouse. Values represent the mean. mice. Compared to control, combining ligand activation of PPARβ/δ with inhibition of COX2 activity caused a delay in the onset of tumor formation in wild-type mice, and this effect was not found in Pparβ/δ-null mice (Figure 4-1A). In response to either GW0742 or nimesulide, the percentage of wild-type mice with skin tumors from week 11 to 16 was lower but not statistically different compared to control wild-type mice (Figure 4-1A). However, in response to both GW0742 and nimesulide, the percentage of wild-type mice with skin tumors from week 11 to 16 was decreased as compared to control wild-type mice (P 0.05; Figure 4-1A). These effects of GW0742, nimesulide or the combination of GW0742 and 202

216 nimesulide were not found in Pparβ/δ-null mice (Figure 4-1A). Skin tumor multiplicity was significantly greater (29-30%) in control Pparβ/δ-null mice as compared to control wild-type mice from week 20 until week 42 of the two-stage bioassay (P 0.05; Figure 4-1B). Ligand activation of PPARβ/δ with GW0742 resulted in decreased (20-40%) tumor multiplicity in wild-type mice during week 37 to week 42 of the bioassay and this effect was not found in Pparβ/δ-null mice (P 0.05; Figure 4-1B). Interestingly, skin tumor multiplicity was lower (24-27%) in Pparβ/δ-null mice in response to topical GW0742 from week 20 to week 30 of the bioassay as compared to control Pparβ/δ-null mice (P 0.05; Figure 4-1B). Dietary nimesulide caused a decrease (30-46%) in tumor multiplicity in wild-type mice during week 24 to week 42 of the bioassay, and this effect was not found in Pparβ/δ-null mice (P 0.05; Figure 4-1B). The combination of topical application of GW0742 and dietary nimesulide resulted in a marked decrease (57-69%) of tumor multiplicity from week 22 onward in wild-type mice and the effect was greater compared to either GW0742 or nimesulide treatment alone from week 21 to week 40 (P 0.05; Figure 4-1B). In Pparβ/δ-null mice, the combination of GW0742 with nimesulide caused a decrease (27-42%) in tumor multiplicity from week 20 to week 30 (P 0.05; Figure 4-1B). Average tumor size was greater in the Pparβ/δ-null mice compared to wild-type mice, but this difference was not statistically significant (Figure 4-1C). Topical GW0742 or the combined treatment of topical GW0742 and dietary nimesulide did not cause a significant decrease of average tumor size in either genotype (Figure 4-1C). Dietary nimesulide caused a decrease in average tumor size in wild-type 203

217 Figure 4-2. Skin tumor size following ligand activation of PPARβ/δ and inhibition of COX2. Wild-type (+/+) and Pparβ/δ-null ( / ) mice were treated with topical GW0742 (5 µm), dietary nimesulide (400 mg/kg) or the combination of GW0742 and nimesulide during a forty-two week two-stage bioassay (initiation with DMBA and promotion with TPA) as described in Methods. A, Incidence of mice with different tumor sizes. These values represent the percentage of mice within a given group that exhibited skin tumors with the indicated size range. B, The distribution of average tumor size for each treatment group. Mice within each treatment were used to calculate the percentage of tumors of that particular size range for each treatment group. *Significantly different than control wild-type, P

218 Figure 4-3. Distribution of keratoacanthomas and squamous cell carcinomas following ligand activation of PPARβ/δ and inhibition of COX2. Wildtype (+/+) and Pparβ/δ-null ( / ) mice were treated with topical GW0742 (5 μm), dietary nimesulide (400 mg/kg) or the combination of GW0742 and nimesulide during a forty-two week two-stage bioassay (initiation with DMBA and promotion with TPA) as described in Methods. Suspected keratoacanthomas and squamous cell carcinomas were examined microscopically and classified as either keratoacanthomas or squamous cell carcinomas by an expert pathologist. Incidence of (A) keratoacanthomas or (C) squamous cell carcinomas. Values represent the percentage of mice with the indicated lesion within each group. Actual number of mice with the indicated lesion within each group of mice is shown in parentheses. Multiplicity of (B) keratoacanthomas or (D) squamous cell carcinomas. Values represent the average number of the indicated lesion per mouse with lesion. 205

219 mice and this effect was not observed in Pparβ/δ-null mice (P 0.05; Figure 4-1C). Closer examination of the distribution of the tumor size also revealed some striking differences (Figure 4-2). The percentage of control Pparβ/δ-null mice with tumors in the 2-3 mm was greater than control wild-type mice (Figure 4-2A). Additionally, the average percentage of total tumors per mouse in the 1-2 mm range was greater in control wild-type mice as compared to control Pparβ/δ-null mice, and this difference was consistent with a greater percentage of total tumors per mouse in the 2-3 mm and greater than 5 mm size ranges in control Pparβ/δ-null mice as compared to control wild-type mice (Figure 4-2B). In wild-type mice fed nimesulide, the percentage of mice with tumors in the 3-5 mm size range, and the percentage of mice with tumors greater than 5 mm in size, was significantly less as compared to control wild-type mice (Figure 4-2A). Similarly, the average percentage of total tumors per mouse greater than 5 mm was lower in wild-type mice fed nimesulide as compared to control wild-type mice (Figure 4-2B). The average percentage of total tumors per mouse in the 1-2, 2-3 and 3-5 mm range was similar in wild-type mice fed nimesulide as compared to control wild-type mice (Figure 4-2B). Dietary nimesulide had no effect on the distribution of tumors with different sizes in Pparβ/δ-null mice as compared to control Pparβ/δ-null mice (Figure 4-2A,B). However, compared to wild-type mice fed nimesulide, the average percentage of total tumors per mouse in the 1-2 mm size range was lower in Pparβ/δ-null mice fed nimesulide (Figure 4-2B). This difference was due to the increase in the average percentage of total tumors per 206

220 mouse in the 2-3 mm and greater than 5 mm size ranges in Pparβ/δ-null mice fed nimesulide as compared to similarly treated wild-type mice (Figure 4-2B). In wild-type mice treated with GW0742, the percentage of mice with tumors in the 2-3 mm size range was greater, while the percentage of mice with tumors in the 3-5 mm and greater than 5 mm size ranges was less as compared to control wild-type mice (Figure 4-2A). This effect was not found in GW0742-treated Pparβ/δ-null mice (Figure 4-2A). GW0742 had no effect on the average size distribution of total tumors per mouse in either genotype (Figure 4-2B). The percentage of wild-type mice treated with both topical GW0742 and dietary nimesulide with tumors in all size ranges was markedly lower as compared to control wild-type mice and this effect was not found in similarly treated Pparβ/δ-null mice (Figure 4-2A). The average percentage of total tumors per mouse in the 2-3 mm range and 3-5 mm ranges was lower in wild-type mice treated with both topical GW0742 and dietary nimesulide as compared to control wild-type mice; these effects were not found in similarly treated Pparβ/δ-null mice (Figure 4-2B). The majority of representative skin lesions examined in all groups were squamous cell papillomas (data not shown). Skin lesions macroscopically suspected of being SCC were examined for histopathology. Skin lesions macroscopically suspected of being SCC were not observed in wild-type mice treated with nimesulide. For control, nimesulide-treated, GW0742-treated and nimesulide+gw0742 treated wild-type mice, 2/8, 0/7, 3/10 and 2/10 mice, respectively, had lesions macroscopically suspected of being SCC. For control, nimesulide-treated, GW0742-treated and nimesulide+gw0742 treated 207

221 Pparβ/δ-null mice, 5/8, 3/10, 4/10 and 5/10 mice, respectively, had lesions macroscopically suspected of being SCC. Histopathological analysis revealed that these lesions were typically either keratoacanthomas or SCC. A higher incidence of keratoacanthoma was observed in control Pparβ/δ-null mice (3/8) compared to control wild-type mice (1/8; Figure 4-3A). No keratoacanthomas were found in wild-type mice fed dietary nimesulide, but neither GW0742, nimesulide or the combined treatment caused any statistically significant changes in the incidence of keratoacanthoma in either genotype (Figure 4-3A). The average number of keratoacanthomas per mouse was comparable between both genotypes, although no keratoacanthomas were noted in wild-type mice fed nimesulide (Figure 4-3B). While 25% of control wild-type mice (2/8) had SCC, no SCC were found in wild-type mice treated with dietary nimesulide or topical GW0742 and only 10% of wild-type mice treated with both dietary nimesulide and topical GW0742 (1/10) had SCC (Figure 4-3C). SCC were found in 25% of control Pparβ/δ-null mice (2/8), 20% of nimesulide-treated Pparβ/δ-null mice (2/10), none of GW0742-treated and 40% of nimesulide and GW0742-treated Pparβ/δ-null mice (4/10) (Figure 4-3C). None of these differences achieved statistical significance. The average number of SCC per mouse was comparable between both genotypes, although no SCC were observed in nimesulide-treated or GW0742-treated wild-type mice or GW0742-treated Pparβ/δ-null mice (Figure 4-3D). One hemangioma was observed in one GW0742-treated Pparβ/δ-null mouse, and one malignant basal cell tumor was found in one nimesulide and GW0742-treated Pparβ/δ-null mouse (data not shown). Interestingly, 208

222 polymorphonuclear neutrophil infiltrates were more commonly observed in Pparβ/δ-null mouse skin lesions as compared to wild-type mouse lesions (Figure 4-4), consistent with past results (Peters et al., 2000b). Additionally, polymorphonuclear neutrophil infiltrates were less common in skin lesions from wild-type mice treated with either nimesulide, GW0742 or the combined treatment, but were more commonly found in similarly treated Pparβ/δ-null mice. 209

223 Figure 4-4. Histopathology of skin tumors following ligand activation of PPARβ/δ and inhibition of COX2. Wild-type (Pparβ/δ+/+) and Pparβ/δ-null (Pparβ/δ / ) mice were treated with topical GW0742 (5 μm), dietarynimesulide (400 mg/kg) or the combination of GW0742 and nimesulide during a forty-two week two-stage bioassay (initiation with DMBA and promotion with TPA) as described in Methods. A, Squamous cell carcinoma (SCC) in control wild-type mouse with polymorphonuclear neutrophils (PMN) infiltration (P); large irregular 210

224 mass invading the dermis (arrows). Magnification = 100X. B, SCC in control Pparβ/δ-null mouse with PMN infiltration (P) and invasion of stroma (arrows). Magnification = 40X. C, SCC in nimesulide-treated Pparβ/δ-null mouse with PMN infiltration (P) and invasion of stroma (arrows). Magnification = 40X. D, Keratoacanthoma in GW0742-treated wild-type mouse. Magnification = 20X. E, Keratoacanthoma in GW0742-treated Pparβ/δ-null mouse with PMN infiltration (P). Magnification = 40X. F, SCC in GW0742 and nimesulide-treated wild-type mouse with invasion of stroma (arrows). Magnification = 100X. G, Malignant basal cell tumor invading adjacent tissue. Magnification = 100X. Bar = 100 μm Effect of GW0742 and nimesulide on terminal differentiation markers Ligand activation of PPARβ/δ or inhibition of COX activity can both induce terminal differentiation in primary keratinocytes and skin (Akunda et al., 2004; Kim et al., 2006a; Schmuth et al., 2004; Tiano et al., 2002; Westergaard et al., 2001). To determine if the enhanced efficacy of inhibiting chemically-induced skin tumorigenesis by combining GW0742 with nimesulide was due in part to modulation of terminal differentiation, expression of differentiation markers was examined. Dietary nimesulide, topical GW0742, and topical GW0742 in combination with dietary nimesulide increased expression of KERATIN 1 (K1) protein in wild-type mouse skin as compared to control, and this effect was not found in Pparβ/δ-null mouse skin (Figure 4-5). Dietary nimesulide or topical GW0742 did not alter expression of KERATIN 10 (K10) protein in mouse skin from either genotype (Figure 4-5). However, topical GW0742 in combination with dietary nimesulide increased expression of K10 protein in wild-type mouse skin as compared to control, and this effect was not found in Pparβ/δ-null mice (Figure 4-5). 211

225 Figure 4-5. Expression of differentiation markers in skin following ligand activation of PPARβ/δ and inhibition of COX2. Protein was isolated from wild-type (Pparβ/δ +/+ ) and Pparβ/δ-null (Pparβ/δ / ) mouse skin following treatment with GW0742, nimesulide or both GW0742 and nimesulide and western blots performed as described in Methods. Hybridization signals for KERATIN 1 (K1) and KERATIN 10 (K10) were normalized to ACTIN. *Significantly different than control wild-type, P Effect of GW0742 and nimesulide on the inflammatory response Inflammation can influence different stages of tumorigenesis. Secretion of pro-inflammatory signaling molecules by immune and somatic cells such as tumor necrosis factor-α (TNFα) and interleukin 6 (IL6) can act on cancer cells and promote tumor growth and malignant conversion (reviewed in (Grivennikov et al., 2010)). The NSAID nimesulide is known to attenuate inflammation by inhibiting COX2 activity and the subsequent production of arachidonic acid metabolites. In addition, ligand activation of PPARβ/δ is also known to have anti-inflammatory activities in rodent and human models (reviewed in (Kilgore and Billin, 2008; Peters and Gonzalez, 2009; Peters et al., 2008)). To determine if attenuation of inflammation could in part underlie the observed inhibition of chemically-induced 212