Cowden Syndrome and Pten Promoter Regulation

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COWDEN SYNDROME AND PTEN PROMOTER REGULATION

DISSERTATION

Presented in Partial Fulfillment of the Requirements for

the Degree Doctor of Philosophy in the Graduate

School of The Ohio State University

Rosemary Elaine Teresi

*****

The Ohio State University 2008

Dissertation Committee: Professor Allen Yates, Advisor Approved by Professor Charis Eng, Co-Advisor

Professor Ching-Shih Chen ______Professor Denis Guttridge Advisor Integrated Biomedical Sciences Professor Matthew Ringel Graduate Program

Professor Kristin Waite

ABSTRACT

Germline mutations of PTEN (phosphatase and tensin homolog deleted on chromosome ten) are associated with the multi-hamartomatous disorder Cowden syndrome (CS). We show here that the PPARγ agonist Rosiglitazone, along with

Lovastatin, Simvastatin, Pravastatin and Fluvastatin can induce PTEN expression by inducing PTEN transcription. Additionally, we observed, for the first time, that upregulation of SREBP protein, known to induce PPARγ expression, can increase PTEN expression. Our results indicate that Rosiglitazone, and SREBP utilizes PPARγ’s transcriptional activity to induce PTEN transcription, while the statins signal through

PPARγ’s protein activity to upregulate PTEN expression. Studing the full-length PTEN identified a region between -854 and -791 that binds an as yet unidentified transcription factor, through which the statins induce PTEN expression.

We examined the downstream effect of five PTEN promoter variants (-861G/T, -

853C/G, -834C/T, -798G/C, and -764G/A) that are not within any known cis-acting regulatory elements. We demonstrated that protein binding to the PTEN promoter (-893 to -755) was not altered in the five variants, when compared to the wild-type (WT) promoter. However, three of the variants (-861G/T, -853C/G, and -764G/A) demonstrated ~50% decrease in luciferase activity compared to the WT construct. PTEN

ii mRNA levels were not altered in these variants, while secondary structure predictions indicated that different PTEN 5’UTR transcript folding patterns exist in three variants, suggesting an inhibition of protein translation. This was confirmed by PTEN protein analysis. These data indicate that variants causing large mRNA secondary structure alterations result in an inhibition of protein translation and a decrease in PTEN protein expression. These data emphasize the importance of PTEN promoter nucleotide variations and their ability to lead to CS progression by a novel regulatory mechanism.

Importantly, these patients have a high prevalence of breast, thyroid and endometrial malignancies, thus understanding of the mechanism of PTEN dysfunction in these patients will lead to more sensitive molecular diagnostic and predictive testing and ultimately, to rational targeted therapies to treat or prevent malignancy.

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DEDICATION

This dissertation is dedicated to the men in my life: Paul Killian, Terry Roof, Stephen Teresi and Rigley Teresi. To my grandfather, Paul, who emphasized the importance of education and always taught me to “keep my eyes on the books and not on the boys”. To my father, Terry, who has been my role model for hard work, persistence and personal sacrifices, and who instilled in me the motivation to set high goals and the confidence to achieve them. To my husband, Stephen, who has been my emotional anchor and provided unending support throughout my doctoral career. Last but not least, to my son Rigley, who is the light of my life.

I would also like to dedicate this work to the rest of my family and friends. You’ve been there for me through both the good and the bad. I will forever be grateful for your continuous encouragement.

ACKNOWLEDGMENTS

My thanks and appreciation to Dr. Charis Eng for persevering with me as my advisor through my time at The Ohio State University and the Cleveland Clinic

Foundation. Her drive and encouragement pushed me to where I am today. I am grateful as well to Drs. Allen Yates and Kristin Waite for coordinating and overseeing my research over the last five years. Additionally to the members of my dissertation committee, Drs. Ching-Shih Chen, Denis Guttridge, Christoph Plass, and Matt Ringel who have generously given their time and expertise to better my work. I thank them all for their many contributions and support.

I would also like to acknowledge all the members of Dr. Eng’s laboratory that

I’ve had the privilege of working with over the last several years; it has been an honor to work with each of you. I would specifically like to acknowledge Drs. Marcus Pezzolesi,

Yufang Tang, Kevin Zbuk, Sarah Planchon-Pope, Guillaume Assié, and Frank Weber, as well as Michelle Sinden, Nita Williams, and Pat Kessler for many helpful discussions over the years. Moreover, I would like to thank Drs. Jodi Bubenik, Donna Driscoll, Don

Luse, and Kwaku Dayie for their insight and Robert Pilarski and Jennifer Stein for aid in patient recruitment.

VITA

July 13, 1980...... Canton, OH

Summer 2001...... Cell Biology Internship University of Cincinnati Cincinnati, OH Mentor: Dr. Karen Knudsen

May 12, 2002...... Bachelor of Science in Biology Bachelor of Science in Chemistry Heidelberg College; Tiffin, OH Mentor: Dr. Robert Murray

2002 – 2005...... Graduate Research Assistant The Ohio State University Columbus, OH Mentor: Dr. Charis Eng

2005 – 2007...... Predoctoral Fellow Cleveland Clinic Foundation Cleveland, OH Mentor: Dr. Charis Eng

PUBLICATIONS

1. Van Brocklyn J.R,, Young N., Roof R. Sphingsine-1-phosphate stimulates motility and invasiveness of human glioblastoma multiforme cells. Cancer Lett 2003 Sep 10; 199(1):53-60.

2. Teresi, R.E., Chaiu, C.W., Chen, C.S., Chatterjee, K.V., Waite, K.A., Eng, C. PTEN’s Transcription Regulated by PPARγ. Int J Cancer 2006 May 15;118(10):2390-8.

vi 3. Weber, F, Teresi, R.E., Broelsch, C.E., Frilling, A, and Eng, C. A Limited Set of Deregulated Human microRNA's Contribute to Follicular Thyroid Carcinogenesis. J Clin Endocrinol Metab 2006 Sep 91(9):3584-91.

4. Teresi R.E., Zbuk K.M., Pezzolesi M.G., Waite K.A., and Eng C. Cowden Syndrome-Affected Patients with PTEN Promoter Mutations Demonstrate Abnormal Protein Translation. Am J Hum Genet. 2007 Oct;81(4):756-67.

5. Teresi R. E., Planchon S.M., Waite K.A., Eng C. Regulation of the PTEN Promoter by Statins and SREBP. Hum Mol Genet, in press (12-04-2007).

6. Teresi R. E., and Waite K.A. PTEN, PPARγ and the Fight Against Cancer. PPAR Research, submitted (02-2008)

FIELDS OF STUDY

Major Field: Integrated Biomedical Sciences

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TABLE OF CONTENTS

P a g e

Abstract...... ii

Dedication ...... iv

Acknowledgments ...... v

Vita ...... vi

List of Tables...... xi

List of Figures ...... xii

List of Abbreviations...... xiv

Chapter 1 INTRODUCTION ...... 1

1.1 History ...... 1

1.2 Cowden Syndrome ...... 1

1.2.1 Phenotype ...... 1 1.2.2 PTEN...... 4

1.3 PTEN Hamartoma Tumor Syndrome ...... 4 1.3.1 Bannayan-Riley-Ravalcaba Syndrome ...... 5 1.3.2 Others ...... 5

1.4 PTEN ...... 6

1.4.1 Mutations...... 6 1.4.2 PTEN Protein Domains...... 8

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1.5 PTEN Functions ...... 10

1.5.1 Nuclear Compartmentalization ...... 10 1.5.2 Phosphatase Activity...... 10

1.6 PTEN Regulation...... 13

1.6.1 Degradation ...... 13 1.6.2 Transcription ...... 14 1.6.2.1 Promoter...... 14 1.6.2.2 Transcription Factors ...... 16 1.6.2.3 Potential PTEN Transcription Inducers ...... 16 1.6.2.3.1 Thiazolidinediones ...... 16 1.6.2.3.2 Statins ...... 17

1.7 Statement of Intent ...... 18

Chapter 2 INCREASED PTEN EXPRESSION DUE TO TRANSCRIPTIONAL ACTIVATION OF PPARγ BY LOVASTATIN AND ROSIGLITAZONE ...... 19

2.1 Introduction ...... 19

2.2 Materials and Methods ...... 22

2.3 Results ...... 27

2.4 Discussion ...... 44

Chapter 3 REGULATION OF THE PTEN PROMOTER BY STATINS AND SREBP .51

3.1 Introduction ...... 51

3.2 Materials and Methods ...... 53

3.3 Results ...... 57

3.4 Discussion ...... 76

ix Chapter 4 COWDEN SYNDROME PATIENTS WITH PTEN PROMOTER MUTATIONS DEMONSTRATE ABNORMAL PROTEIN ...... 80

4.1 Introduction ...... 80

4.2 Materials and Methods ...... 82

4.3 Results ...... 88

4.4 Discussion ...... 104

Chapter 5 DISCUSSION ...... 111

REFERENCES ...... 118

LIST OF TABLES

Table Page

1.1 Current Operational Diagnostic Criteria for the Diagnosis of CS...... 3

4.1 Increase cancer frequency in Cowden syndrome patients with PTEN promoter variants ...... 88

LIST OF FIGURES

Figure Page

1.1 PTEN Mutations...... 7

1.2 PTEN Domains...... 8

1.3 PTEN Phosphatase Activity ...... 11

1.4 PTEN Promoter ...... 14

2.1 Lovastatin and Rosiglitazone induce PTEN protein expression in a dose- dependent manner...... 28

2.2 Rosiglitazone’s analog, Compound-66, has a minimal effect on PTEN protein expression ...... 39

2.3 Lovastatin and Rosiglitazone inhibit P-AKT and P-MAPK protein expression ...... 31 2.4 Lovastatin and Rosiglitazone, but not Compound-66 induce G1 arrest ..33

2.5 Lovastatin and Rosiglitazone induce PTEN mRNA expression ...... 35

2.6 Lovastatin and Rosiglitazone induce PPRE regulated transcription...... 38

2.7 Upregulation of PTEN expression due to Lovastatin and Rosiglitazone is PPARγ-dependent and not cell line specific ...... 42

3.1 Simvastatin, Pravastatin and Fluvastatin induce PTEN protein expression in a dose-dependent manner ...... 58

3.2 Simvastatin, Pravastatin and Fluvastatin induce PTEN mRNA expression and PPRE regulated transcription ...... 59

3.3 Three potential models of statin-mediated PTEN upregulation ...... 62 xii

3.4 PTEN upregulation by statins and ALLN are not cell line-specific...... 64

3.5 SREBP induces PPARγ-mediated transcription, which is inhibited by the statins...... 67

3.6 SREBP signaling not inhibited by Rosiglitazone ...... 70

3.7 Statins induce PTEN transcription within the minimal promoter...... 72

4.1 Schematic diagram of the PTEN promoter ...... 89

4.2 EMSA analysis probed with radiolabelled PTEN promoter from -893 to - 755, which is either WT or variant ...... 92

4.3 Luciferase activity of PTEN 5’UTR altered in three VUSs ...... 94

4.4 WT and variant PTEN promoters demonstrate equal mRNA expression 97

4.5 MFOLD-predicted secondary structures resulting from the five VUS in Cowden syndrome patients ...... 100

4.6 PTEN protein expression decreased in promoter variants with the greatest mRNA secondary structure alterations ...... 102

5.1 -764G/A PTEN Promoter Mutation Produces a Novel Transcription Factor Binding Site...... 116

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LIST OF ABBREVIATIONS

CS Cowden syndrome

BRRS Bannayan-Riley-Ravalcaba syndrome

Cmpd-66 Compound-66

Egr-1 Early growth response-1 eIF4F Eukaryotic translation initiation factor

EtOH Ethanol

LBCL lymphoblastoid cell line

Lov Lovastatin

MAPK Mitogen-activated protein kinase

MEF Mouse Embryonic Fibroblast

PHTS PTEN harmatoma tumor syndromes

PIP3 Phosphatidylinositol (3,4,5) triphosphate

PIP2 Phophatidylinositol (4,5) biphosphate

PTEN Protein phosphatase and tensin homolog located on chromosome ten

Rosi Rosiglitazone

NaBut Sodium Butyrate

TZD Thiazolidinediones

VUS Variant of unknown significance

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CHAPTER 1

INTRODUCTION

1.1 History

In 1963 Lloyd and Dennis presented a report on a previously undefined disorder,

Cowden syndrome (CS), which has since changed the way we look at cancer. They

described a patient, Rachel Cowden, who presented with multiple mucocutaneous

abnormalities whom later died of infiltrating ductal carcinoma of the breast at 30 years of

age(1). This observation has provided the necessary foundation for understanding cancer

development as well as point scientists down the path to personalized health care.

1.2 Cowden Syndrome

1.2.1 Phenotype

CS is an autosomal dominant disorder characterized by multiple haramatomas of the three germ layers and increased risk of neoplasia. This syndrome affects approximately 1 in 200,000 individuals, where 10% to 50% of cases are thought to be familial in nature(2). However, the incidence of CS cases is generally thought to be an underestimate due to its complex variability(3, 4). In 1995, the International Cowden

Consortium (ICC) set a uniform diagnostic criterion to properly identify these patients(3,

5), which was subsequently updated in 2000(2). The ICC has characterized CS by five major criteria including breast carcinoma, thyroid carcinoma, macrocephaly, Lhermitte-

Duclos disease (LDD) and endometrial carcinoma (Table 1.1)(2). Patients diagnosed with

CS have a 25-50% lifetime risk of developing female breast cancer, while the risk for the general population is ~13%. Currently an increased risk of male breast cancer is debatable(3). Moreover, breast cancer in CS patients is also thought to express ~10 years prior the general population(7, 8). CS patients also have an increased risk for epithelial thyroid cancer, which tends to be follicular. Patients have ~10% lifetime risk of developing thyroid cancer, compared to <1% in the general population(7-9).

Furthermore, there is a ~5-10% lifetime risk of endometrial cancer in CS patients compared to ~2-4% in the general population(6, 10). Additionally, CS is characterized by seven minor criteria, which in combination with the above major symptoms can indicate disease (Table 1.1). Approximately 90% of CS patients will demonstrate a phenotype by the age of 20, while 99% will observe signs of mucocutaneous lesions by 29 years of age(2, 3). Proper recognition and diagnosis of CS patients is crucial not only due to the increased risk of cancers observed in this syndrome, but also due to the morbidity and even mortality associated with its non-malignant features.

Table 1.1 Current Operational Diagnostic Criteria for the Diagnosis of CS(2)

Pathognomonic Criteria Operational Diagnosis in an Individual Adult Lhermitte-Duclos disease (LDD) Mucocutaneous lesions alone if: Mucocutaneous lesions Six or more facial papules, of which >3 must Trichilemmomas, facial be trichilemmomas, or Cutaneous facial Acral keratoses papules and oral mucosal papillomatosis, or Papillomatous papules Oral mucosal papillomatosis and acral Mucosal lesions keratoses, or Palmoplantar keratoses, >6

>2 major criteria or One major and >3 minor or >4 minor

Major Criteria: Operational Diagnosis in a Family Where Breast cancer One individual is diagnostic for Cowden Thyroid cancer (esp. follicular) syndrome: Macrocephaly (megalencephaly) (ie, The pathognomonic criteria >97th percentile) Any 1 major criteria with or without minor Endometrial cancer criteria History of BRRS Two minor criteria

Minor Criteria: Other thyroid lesions (eg, multinodular goiter, adenoma) Mental retardation (ie, IQ <75) Gastrointestinal hamartomas Fibrocystic breast disease Lipomas Fibromas Genitourinary tumors (especially renal cell carcinoma) Genitourinary manifestations Uterine fibroids

1.2.2 PTEN

In 1996, our laboratory mapped the CS susceptibility gene to 10q22-23 by studying twelve families with classic CS(3). Utilizing the meticulously obtained localization of the critical interval allowed two groups to identify an important tumour suppressor, PTEN/MMAC1 in that region. Steck and colleagues named their gene phosphatase and tensin homolog deleted on chromosome ten (PTEN)(11), while Li et al coined the term mutated in multiple advanced cancers (MMAC1)(12). Independently while looking for TGF-β responsive phosphatase molecules, Hong Sun’s group identified transforming growth factor regulated and epithelial enriched phosphatase (TEP1), which turns out to be identical to PTEN(13). Currently PTEN is the generally accepted GDB name. More detailed analysis of PTEN indicate that it is a widely expressed nine exon gene which composes an ~5.5 kb mRNA and 403 amino acid protein. Our laboratory was the first to identify germline PTEN mutations in 4 of 5 CS families, thus establishing

PTEN as the CS gene(4). Because of careful genetic and clinical studies, our laboratory has established an approximately 85% mutation frequency in those with classic CS(5,

6)(14).

1.3 PTEN Hamartoma Tumor Syndrome

After the identification of PTEN as the susceptibility gene for CS, similar features between CS and some of the other hamartomatous syndromes led our laboratory to explore PTEN as a susceptibility gene for these seemingly clinically disparate syndromes.

This led to the identification of germline PTEN mutations in a subset of patients within these additional hamartomatous syndromes, thus grouping these syndromes together as

PTEN hamartoma tumor syndromes (PHTS). PHTSs encompasses PTEN mutation positive CS, as described above, Bannayan-Riley-Ravalcaba syndrome (BRRS [MIM

153480]), Proteus syndrome (PS [MIM 176920]), Proteus-like syndrome (PSL), autism, and VATER (7).

1.3.1 Bannayan-Riley-Ravalcaba-Syndrome

Germline PTEN mutations have been found to be associated with BRRS. This disorder is characterized by macrocephaly, lipomatosis, hemangiomatosis, developmental delay and speckled penis(8), however formal diagnostic criteria have yet to be established. It is currently thought that 65% of BBRS patients have a germline PTEN mutation(14, 17). Prior to the identification of PTEN mutations, BRRS was not thought to be associated with increased cancer risk. However, with the demonstration of PTEN mutations, the recommendation is that BRRS patients follow the same surveillance and management guidelines as CS patients.

1.3.2 Others

PS and PSL are complex disorders that affect multiple organs which is characterized by congenital malformations, hemihypertrophy, hamartomatous overgrowths, epidermal nevi and hyperostosis(9). This multi-organ syndrome suggests that it may be inherited through a mosaic germline mutation. Additionally, PS and PSL can also occur sporadically, suggesting that the mutation may be somatic. Indeed a PTEN germline mosaic mutation was uncovered in a PSL patient(10) and somatic PTEN nucleotide alterations have also been observed(11). Another PHTS is a subset of autism

spectrum disorders. Our laboratory was the first to establish that 17% of patients with

autism spectrum disorder who also havemacrocephaly harbored germline PTEN

mutations(21). Finally, a germline PTEN mutation has also been identified in an

individual with VATER, which also included the features of megalencephaly and autism.

VATER is characterized by vertebral and anal malformations, tracheoesophageal atresia,

and radial and renal malformations(22). The above disorders are all considered to be

PHTS and with the demonstration of PTEN mutations, the recommendation is that these

patients follow the same surveillance and management guidelines as CS patients.

1.4 PTEN

1.4.1 Germline Mutations

85% of CS and 65% of BBRS patients have an identified PTEN mutation.

Interestingly, the location of PTEN mutations appears to be interchangeable between CS

and BRRS patients indicating that they are allelic (Figure 1.1). Theses studies have

demonstrated that ~70% of PTEN mutations lie within exons 5, 7 and 8. Moreover, 40%

of all PTEN mutations lie within exon 5, despite it only representing 20% of its coding

region(23). Groups have tried to perform genotype-phenotype correlation; however

results were inconclusive. One study demonstrated PTEN germline mutation families

have an increased risk of being diagnosed with malignant breast cancer, compared to

those that do not have an identified mutation. Moreover, they were also able to show that

mutations located within the 5’ end of exon 5 and all PTEN missense mutations were associated with multi-organ involvement, hence a more severe phenotype(23).

Figure 1.1 PTEN Mutations(7)

PTEN mutations identified within CS (circles), BRRS (squares), CS-BRRS overlap

(diamonds), PS (hearts) and PSL (crosses) patients.

1.4.2 PTEN Protein Domains

PTEN’s protein product, PTEN, is composed of an N-terminal and a C-terminal domain (Figure 1.1). Amino acids 1-185 comprise the N-terminal domain, while amino acids 186-403 encompass the C-terminal domain. Interstingly, 57% of all PTEN

mutations occur in the N-terminus, while the remaining 43% occur within the C-

terminus(7).

The N-terminus of PTEN contains the phosphatase domain (Figure 1.2).

Approximately 31% of all PTEN mutations lie within the active phosphatase domain, 7

thus demonstrating its biological importance. Interestingly this domain is a slightly larger

then most active phosphatase domains. This allows for both proteins and phospholipids to

be substrates therefore making PTEN a unique phosphatase(24).

Figure 1.2 PTEN Domains(7)

PTEN is composed of a N-terminus which contains the phosphatase domain the C- terminus. PTEN’s C-terminus is composed of the C2 (blue box), PEST (orange striped boxes) and PDZ (red box) domains. PTEN can be phosphorylated (blue asterisks) within the PEST domain in the C-terminus.

The C-terminus of PTEN contains three major domains. The domains from 5’ to 3’ are

C2, PEST, and PDZ (Figure 1.2). These three domains are involved in PTEN signaling, stability, and necessary for proper function. The role of the C2 domain is to interact with phospholipid membranes, thus allowing PTEN to function at the plasma membrane(24). 8

The C2 domain additionally allows for PTEN to network with proteins implicated in signal transduction and membrane localization(12). Further 3’ is the PEST domain, which is important for PTEN stability. Georgescu and colleagues showed that deletion of this domain caused a decrease of PTEN protein expression(13). The last domain in the C- terminus is the PDZ domain, which is necessary for protein-protein interaction. Similar to the C2 domain, these interactions are important for signaling transduction(27, 28).

1.5 PTEN Functions

1.5.1 Nuclear Compartmentalization

Prior to 1998, PTEN was thought to only function in the cytoplasm. However, recently, it has been shown to be located in the nucleus of the cell as well (29).

Interestingly, immunohistochemistry on thyroid tissue demonstrated a higher concentration of PTEN in the nucleus of normal tissue when compared to malignant tissue(30). These data indicate that the cellular location of PTEN is significant in a cancer setting. PTEN does not have a traditional nuclear localization sequence thus making it difficult to determine how it translocates to the nucleus. Recently our laboratory has implicated major vault protein (MVP) in moving PTEN across the nuclear membrane, at least in breast cancer cells(31, 32). However, the exact mechanism has yet to be established. Moreover, the precise function of nuclear PTEN has yet to be concretely determined, but research has shown it to play a pivotal role in the cell cycle, more specifically, G1 arrest(33). We have shown that there is an increase in the localization of

PTEN to the nuclear compartment when cells are in G0-G1 of the cell cycle(14). PTEN

has also been associated with the regulation of pro-apoptotic transcription factors in the nucleus. More recently PTEN has been implicated in associating with centrimeres, indicating that it plays a role in chromosomal stability.

1.5.2 Phosphatase Activity

As stated above, PTEN is a unique phosphatase that has the ability to dephosphorylate both proteins and lipids (Figure 1.3). Its lipid phosphatase activity functions as a negative regulator of AKT phosphorylation. PTEN dephosphorylates phosphatidylinositol-(3,4,5)-triphosphate (PIP3) at the D3 position generating phosphatidylinositol-(4,5)-biphosphate (PIP2), subsequently decreasing PIP3 levels.

Since PIP3 is required for AKT phosphorylation, active PTEN leads to a decrease in P-

AKT levels and consequently a decrease in AKT-mediated proliferation pathways(38,

39). The protein phosphatase activity of PTEN has been shown to inhibit the

Shc/Grb2/Sos and mitogen-activated protein kinase (MAPK) pathways. The dephosphorylation of Shc by PTEN indirectly decreases the phosphorylated form of

MAPK levels, thus reducing MAPK’s activity(40). Additionally, PTEN’s protein phosphatase activity up-regulates p27 with a concomitant down regulation of cyclin D1 thus coordinating G1 arrest(33, 41). By regulating these key signaling pathways, PTEN downregulates cell division and upregulates apoptosis. Additionally, PTEN’s protein phosphatase activity has been shown to dephosphorylate focal adhesion kinase (FAK), which inhibits cell spreading and migration(29, 42).

Figure 1.3 PTEN Phosphatase Activity(15)

PTEN’s lipid (left pathway) and protein (right pathway) phosphatase activity regulates several signaling pathways. PTEN’s lipid phosphatase activity regulates the phosphoylation of AKT, while its protein phosphatase activity regulates the phosphorylation of FAK and MAPK.

1.6 PTEN Regulation

1.6.1 Degradation

Within the last 50 amino acids of PTEN’s C-terminus are five phosphorylation sites on serine residues 370, 380, and 385 and threonine residues 382, and 383(44). Of these sites the protein kinase, casein kinase 2 (CK2), phophorylates four of them: S370,

S380, T383 and S385(16). Research indicates that PTEN phosphorylation is involved in its protein stability, but does not play a role in its normal function. More specifically, studies have shown that S380, T382 and T383 are crucial for PTEN protein stability.

When these sites are mutated, PTEN’s protein half-live is decreased and subsequently is its protein levels. Data has suggested that these PTEN levels are decreased through proteasome-mediated degradation(16). Recently, PTEN phosphorylation has also been shown to cause a conformational change that masks the PDZ domain, reducing PTEN’s ability to bind to PDZ-domain containing proteins(44).

In 2006, our laboratory demonstrated that p53 has the ability to regulate PTEN

degradation as well(17, 18). By using p53–/– MEF and double PTEN–/–, TP53–/– PC3

cells, we were able to reveal that the stabilization and activation of p53 through

proteasome inhibition leads to down-regulation of PTEN. These works indicate that the

induction of p53 leads to an increase in proteasome-mediated degradation of PTEN,

presumably through the caspase pathway. This significant study coorelates with several

early stage primary cancer clinical results; where a decrease in PTEN protein expression

is observed.

1.6.2 Transcription

1.6.2.1 Promoter

The promoter region of PTEN was initially characterized by Sheng and co- workers in 2002 (Figure 1.4)(19). In their study, they mapped PTEN’s minimal promoter to nucleotide positions -958 to -821 and the core promoter to -1344 to -745, where +1 signifies the ATG site. The location of PTEN’s transcriptional start site has yet to be

concretely established; however several laboratories have suggested a number of putative

start sites(46-49). The majority of these sites lie between -1031 and -93; however

comparison of the human and mouse PTEN cDNA sequences suggest that its transcript

begins around -925(20). PTEN is therefore unique in that its promoter and 5’ UTR

overlap.

Prior to 2003, PTEN mutations had been identified in 80% of patients meeting

strict diagnostic criteria for CS(5). This syndrome is believed to be mono-genic, therefore

determining the cause of PTEN dysfunction in the remaining 20% of patients is vitally

important to the practice of personalized genetic healthcare. Our laboratory began

aggressively pursuing alternative mechanisms of PTEN inactivation including

interrogating the mutation status of its own promoter. We have since identified mutations

in the PTEN promoter within CS patients and have shown that ~10% of previously

classified PTEN mutation negative patients actually have nucleotide variants within the

full-length promoter. Furthermore, these mutations resulted in both a decrease in PTEN

protein expression and loss-of-functional activity. Interestingly, in this initial study, eight

of nine (89%) patients with germline PTEN promoter mutations had breast cancer,

although the overall number of organs involved was generally low in these patients

suggesting these mutations may preferentially confer very high penetrance for breast

cancer(14).

Figure 1.4 PTEN Promoter(50)

PTEN’s full-length promoter lies between -1344 and -745 (white bar), while the minimal promoter lies between -958 and -821 (gray bar). Four transcription factors are thought to lie within the promoter: CBF-1 (blocked bar), p53 (dashed bar), EGR1 (dotted bar) and

Sp1 (black bar). 14

1.6.2.2 Transcription Factors

Transcriptional regulation of PTEN is beginning to be elucidated, however, there is still much to be understood. To date, analysis of PTEN’s promoter has identified eight regulatory factors that have been implicated in modulating PTEN’s transcription (Figure

1.4). In 2001, Stambolic et al identified a functional p53 binding site required for its upregulation, located at nucleotide positions -1190 to -1157 in PTEN’s promoter(20).

Additionally, early growth response-1 (Egr-1) has been shown to bind to positions -947 to -939 to induce PTEN expression(51). In 2001, two putative binding sites for the transcription factor peroxisome proliferator-activated receptor-gamma (PPARγ) were identified approximately 10kb upstream of the minimal promoter region of PTEN(21); specific binding of PPARγ was later confirmed(22). Recently our laboratory has identified a USF1 binding site ~2 kb (-2237 and -2058) upstream of the ATG site(23).

CBF-1(24) and c-Jun(25) have also recently been indicated as PTEN transcription factors.

More recently, suppression of PTEN gene expression has been shown by the tumor necrosis factor-alpha/nuclear factor-kappaB (NF-κB)(26). The precise mechanism of this inhibition, however, remains unclear.

1.6.2.3 Potential PTEN Transcription Inducers

1.6.2.3.1 Thiazolidinediones

The two PPARγ binding sites upstream of PTEN’s minimal promoter region(21), suggested that PPARγ, acting as a tumor suppressor in breast cancer, can upregulate the transcription of PTEN. These results warranted a full investigation of PPARγ-mediated 15

transcription of PTEN at the biochemical and molecular level; more specifically the

PPARγ agonists ability to induce PTEN expression(57). The clinically used PPARγ agonists are known as thiazolidinediones (TZD)(58). Four recognized TZDs are

Ciglitazone (Alexis), Pioglitazone (Actos), Rosiglitazone (Avandia) and Troglitazone

(Rezulin). PPARγ plays a key role in lipid metabolism, therefore the TZDs are clinically used to treat type II diabetes mellitus.

1.6.2.3.2 Statins

The statin family of drugs, similar to the TZDs, can downregulate lipid metabolism, yet the statins major target is 3-hydroxy-3-methyl-glutaryl-CoA reductase

(HMG-CoA). However, due to the statins’ role in lipid regulation(27), they were also hypothesized to potentially act as a PPARγ agonist. This suggests that the statins may play a key role in inducing PTEN expression. Mevastatin was the first HMG-CoA reductase inhibitor isolated in 1971 in Penicillim citrinum by Dr. Akira Endo. By the mid-1980’s it was being studied clinically to aid patients with hypercholesterolemia. In

1978 Lovastatin was isolated from Aspergillus terreus, and became approved by the Food and Drug Administration (FDA) in 1987 under the name Mevacor. Altering natural statins or creating synthetic compounds subsequently lead to the development of a number of other statins including Simvastatin (Zocor), Pravastatin (Livals), and

Fluvastatin (Lescol)(28). Currently statins are used within millions of patients to lower cholesterol levels and prevent cardiovascular disease.

1.7 Statement of Intent

These previous studies led us to hypothesize that the regulation of PTEN

transcription is key to maintaining the appropriate levels of PTEN needed to combat

cancer development. Our initial studies investigated if there are any potential clinical

PTEN transcription inducers, such as the TZDs and statins. Indeed we demonstrate that

Rosiglitazone, Lovastatin, Simvastatin, Pravastatin and Fluvastatin can induced PTEN

expression by upregulating its transcription. However, our results indicate that the

mechanism utilized by Rosiglitazone is separate from the four statins. Moreover, through

PTEN promoter analysis, we further demonstrated that the statins upregulate PTEN

transcription through an unknown transcription factor(s) located between -893 and -755.

Interestingly, in an independent study we observed that CS mutations located within this

region lead to a decrease in PTEN protein expression due to protein translation inhibition.

In conclusion, these results point to the importance of the PTEN promoter in CS development.

CHAPTER 2

INCREASED PTEN EXPRESSION DUE TO TRANSCRIPTIONAL ACTIVATION OF

PPARγ BY LOVASTATIN AND ROSIGLITAZONE

2.1 Introduction

Breast cancer is the second leading natural cause of death in women and is thought to affect over 13% of women in the United States(29). Germline mutations in

PTEN, a tumor suppressor gene, are associated with 85% of patients with the autosomal dominant disorder CS(14, 17, 64). Patients with CS have a 25%-50% risk of developing breast cancer(65, 66). CS patients have also been reported to have mucocutaneous lesions, thyroid abnormalities, fibrocystic disease, uterine leiomyoma and macrocephaly(2, 67). Germline PTEN mutations are also associated with 65% of patients with BRRS, which is characterized by macrocephaly, lipomatosis, hemangiomatosis and speckled penis(14, 16). Somatic alterations in PTEN, whether by genetic or epigenetic mechanisms, play some role in the pathogenesis of a broad range of solid tumors, such as carcinomas of the breast, thyroid, endometrium and colon(30).

PTEN’s protein product, PTEN, is a dual specificity phosphatase with both lipid and protein phosphatase activity(31). Its lipid phosphatase activity functions as a negative

regulator of AKT phosphorylation. PTEN dephosphorylates PIP3 at the D3 position

generating PIP2, subsequently decreasing PIP3 levels. Since PIP3 is required for AKT

phosphorylation, active PTEN leads to a decrease in P-AKT levels and consequently a

decrease in AKT-mediated proliferation pathways(32). The protein phosphatase activity

of PTEN has been shown to inhibit the Shc/Grb2/Sos and MAPK pathways. The

dephosphorylation of Shc by PTEN indirectly decreases the phosphorylated form of

MAPK levels, thus reducing MAPK’s activity(42). By reducing the cellular levels of both

P-AKT and P-MAPK, PTEN downregulates cell division and upregulates apoptosis.

Transcriptional regulation of PTEN is beginning to be elucidated, however, there is still much to be understood. Several groups have shown that PTEN transcription may be upregulated by Egr-1(51), p53(20), and Sp1(33). In contrast, NF-κB has recently been shown to inhibit PTEN transcription by a yet undetermined mechanism(26). In 2001, two putative binding sites for the transcription factor PPARγ were identified in the 10kb upstream of the minimal promoter region of PTEN(21). However, the biological importance of these sites and what role PPARγ plays in PTEN transcription has not been determined, and so it remains to be determined if PPARγ has a role in PTEN transcription, and if so, which regions of the PTEN promoter are responsible. PPARγ is a transcription factor that regulates gene expression by binding to PPARγ response elements (PPRE) within a promoter of a target gene. Until recently, the focus of PPARγ

research has been on its role in regulating fatty acid metabolism. However, recent

evidence indicates that PPARγ may also act as a tumor suppressor by mediating the

transcription of genes necessary for anti-proliferation and pro-differentiation, which

ultimately can downregulate cellular growth and upregulate apoptosis(17, 18). Increasing

evidence has also shown that PPARγ activation can be anti-carcinogenic, indicating the

treatment potential for PPARγ agonists in several diseases(71, 72). Furthermore, the

addition of PPARγ agonists to breast cancer cell lines have been shown to cause them to

differentiate(73). Previously, we have shown that somatic loss-of-function mutations in

PPARγ plays a role in colorectal carcinogenesis(74) and that PPARγ is frequently downregulated in follicular thyroid carcinomas(34). Additionally, PPARγ has been

shown to be under-expressed and correlates with clinical outcomes in breast cancer

patients(76). In nude mouse and rat models, treatment of breast cancer cells with PPARγ

agonists inhibits tumor progression within the breast tissue(77, 78). There is also an

overlap of solid tumor data where both PPARγ and PTEN have been implicated in its

pathogenesis(79, 80). Thus, in vivo and in vitro evidence is mounting to support the idea

of PPARγ as a tumor suppressor.

Based on these data, we hypothesize that PPARγ, acting as a tumor suppressor in breast cancer, can upregulate the transcription of PTEN and believe that it is warranted to fully investigate PPARγ-mediated transcription of PTEN at the biochemical and molecular level. In order to investigate the effect of PPARγ agonists on PTEN expression, we examined the ability of five PPARγ agonists that have been or are routinely used in the clinical setting, to induce PTEN expression in the MCF-7 breast cancer cell line. Four of the agonists, Ciglitazone, Pioglitazone, Rosiglitazone (Rosi) and

Troglitazone, are TZD and are used to treat type II diabetes mellitus, due to their ability to directly activate PPARγ(57, 58). Currently Lovastatin is used clinically for its statin

activity of regulating high cholesterol levels. While Lovastatin is generally believed to be

a PPARγ agonist, no group has analyzed its ability to directly activate PPARγ. We will

show here, substantive data that indicate that Lovastatin and Rosi induce PTEN protein

via a PPARγ-mediated mechanism.

2.2 Materials and Methods

Materials

Ciglitazone was obtained from BIOMOL Research Laboratories Inc (Plymouth

Meeting, PA). Rosiglitazone was a generous gift from Dr. Lisa Yee (The Ohio State

University). Both Pioglitazone and Troglitazone were obtained from Cayman Chemical

(Ann Arbor, MI). Lovastatin and Cyclohexamide were purchased from Sigma-Aldrich

(St. Louis, MO). Antibodies were obtained from: Cascade Bioscience, Waltham, MA

(PTEN 6H2.1) and Cell Signaling (P-p44/42, P-AKT S473, AKT, and Actin; Beverly,

MA). Cell culture media was obtained from Gibco-BRL (Rockville, MD). M-PER

Mammalian Protein Extraction Reagent was obtained from Pierce Biotechnology Inc.

(Rockford, IL). All other reagents were purchased from standard commercial sources.

Cell culture and stimulation

The MCF-7 breast cancer cell line was maintained at 37°C with 5% CO2 in

DMEM containing 10% FBS and 100 units/ml each penicillin and streptomycin. Both the MEF-WT and the MEF-PPARγ null cell lines were maintained at 37°C with 5% CO2 in DMEM containing 14% FBS and 100 units/ml each penicillin and streptomycin. BT- 21

549 cells were maintained at 37°C with 5% CO2 in DMEM containing 10% FBS and insulin. Cells were plated at 1.0 x 108 24 hrs prior to treatment. After 24 hrs, stimulates dissolved in EtOH, were added as indicated in figure legends. The cells were incubated for an additional 48 hrs or as otherwise indicated in the figure legends. In experiments stimulated with CHX, 10 µg/ml CHX was incubated with MCF-7 cells for either 24 or 48 hrs, as indicated. In experiments stimulated with NaBut, 2.5 mM was incubated with

MCF-7 cells for 48 hrs.

Synthesis of PPARγ Analogs

Synthesis of the following analogs were performed as previous described (57):

∆TG (5-[4-(6-hydroxy-2.5,7,8-tetramethyl-chroman-2-ylmethoxy)-benzylidene]-2,4- thiazonlidine-dione), ∆CG (5-[4-(1methyl-cyclohexylmethoxy)-benzylidene]- thiazonlidine-2,4-dione), Cmpd-66 (5-[4-(2-methyl-pyridin-2-yl-amino)-ethoxyl]- benzylidene]- thiazonlidine-2,4-dione), and ∆PG (5-[4-(2-[5-ethyl-pyridin-2-yl]-ethoxyl)- benzylidene]-thiazolidine-2.4-dione). The identity and purity (>99%) of these TZD derivatives were determined by proton magnetic resonance, high-resolution mass spectrometry and elemental analysis.

Protein Extraction

MCF-7 cells were plated and stimulated as described above. At the time of harvest, media was removed and the cells were washed with PBS. Cells were then harvested into M-PER lysis buffer containing PMSF (0.75mg/ml), benzamidine hydrochloride (0.5 mg/ml), leupeptin (2µg/ml), aprotinin (2 µg/ml), pepstatin (2 µg/ml), 22

β-glycerophosphate (10 mM), NaOV (0.2 mM), and NaF (25mM). Cells were incubated

at room temperature with lysis buffer for 1 min before harvesting. Samples were then

centrifuged at 16,000 X g for 10 min at 4ºC to remove cellular debris. The resulting

supernatant was stored at -80°C. Protein concentration was determined using the bicotinic method(35) using BSA as a standard.

Western Blot

Proteins (30µg) were prepared by the Laemelli method(36), then separated on a

10% SDS-PAGE gel, and electrophoretically transferred onto nitrocellulose(37). Equal

protein loading between conditions was confirmed by staining with Ponceau S solution.

Non-specific binding was blocked by incubating the nitrocellulose blots with 5% milk in

TBS-T (100mM Tris, pH, 1M NaCL, 1% Tween-20) for 1 hr at room temperature. Blots

were then incubated with the primary antibody (1:1000 in 3% BSA) for 2 hrs at room

temperature. Following the primary incubation the blots were washed with TBS-T for 1

hr with frequent changes of buffer. Blots were then incubated with the appropriate

secondary antibody conjugated to horseradish peroxidase (Promega; Madison, WI)

(1:2500 dilution in 5% milk) overnight at 4°C, and washed with TBS-T for 1 hr. Protein

bands were visualized using enhanced chemiluminescence as described by the

manufacturer (Amersham Pharmacia Corp; Piscataway, NJ). The resultant films were

then quantified using NIH-Imager densitometry software.

RT-PCR and Real-Time PCR

MCF-7 cells were stimulated as described above. After stimulation, cells were released by trypsin treatment and subsequently washed three times with PBS through centrifugation. RNA was extracted from the cells following the Qiagen RNeasy Mini

Protocol and then converted to cDNA by Superscript II Reverse Transcriptase (SSIIRT).

The resultant cDNA was subjected to multiplex PCR amplification using primers specific to exon 1 and exon 9 of PTEN (5’ TCAAGAGGATGGATTCGACTT 3’; 5’

TGAAGTACAGCTTCACCTTAAA 3’) and primers to B-actin (Quantum RNA β-actin;

Ambion INC; Austin, TX). Primers were allowed to anneal at 58°C for 30 seconds. The products from the PCR reactions were run on a 1% agarose gel containing ethidum bromide and visualized under a UV light. The real-time quantitative PCR and analysis were carried out using the ABI 7700 Sequence Detector System (ABI/Perkin Elmer,

Foster City, CA) as previously described (14). PTEN exons 1 and 5 were chosen as targets for the real-time quantitative PCR assay. The primers and probes were designed by using PRIMER EXPRESS software (Applied Biosystems). The PTEN exon 3 and exon 5 probes were labeled at the 5’ ends with the reporter dye FAM and at the 3’ end with the MGBNF quencher. GAPDH was used as an internal control. The real-time quantitative multiplex PCR assay was optimized according to the instructions in User

Bulletin No.5. The thermal cycling conditions were 50oC for 2 min and 95oC for 10 min, followed by 40 cycles of denaturing at 95oC for 15 s and annealing and extension at 60oC for 1 min.

Cell Cycle Assay

MCF-7, MET-WT, MEF-PPARγ null and BT-549 cells were plated and

stimulated as described above, after stimulation cells were harvested through

trypsinization. 1.0 x 106 were then resuspended in 70% EtOH and stored at -20°C until

analysis. Cells were stained with 1 µg/ml PI in PBS containing 0.1% Triton X-100. Flow

cytometry was performed on a Beckman-Coulter elite flow cytometer using a 610 long

pass filter for data collection. Data were filtered, and cell cycle phases were quantified using the Modfit program (Verity Software, Bowdoin, ME)(14).

Reporter Assay

Plasmids were co-transfected into 6-well cultures of MCF-7 cells in serum-free media with DMRIE-C (Invitrogen). Each well was co-transfected with 500 ng

PPRETKLUC, 50 ng Renilla luciferase control plasmid and 100 ng of receptor expression vector (pcDNA3, WT PPARγ1, or L468A/E471A PPARγ1). Each plasmid has

previously been described(84). After 12 hr the cultures were washed and new media

containing EtOH, 30 µM Rosi, 3 µM Cmpd-66 or 3 µM Lovastatin was added for 48 hr.

Cells were then harvested using luciferase lysis buffer as described by the manufacturer

(Promega). Samples were analyzed on a luminometer using Renilla luciferase as an internal transfection control.

Statistical Analysis

Statistical analysis was done using Student’s t test. Data are mean ± SD of three

independent experiments and normalized to a control. P<0.05 is considered significant. 25

2.3 Results

Lovastatin and Rosiglitazone (Rosi) induce PTEN protein expression in a dose- and time-

dependent manner

To determine PPARγ’s role in regulating PTEN expression, we examined five

PPARγ agonists’ ability to induce PTEN expression in MCF-7 breast cancer cells. We

observed that stimulation with Ciglitazone, Pioglitazone and Troglitazone did not alter

PTEN expression, despite a wide range of concentrations tested (0-100 µM), but they did

inhibit cyclin D1 in a dose-dependent manner as previously described (data not

shown)(85). Prior studies, which tested Rosi’s ability to induce PTEN in MCF-7 cells,

studied its effect at only one dose, 1µM (21). Studying Rosi’s effects on PTEN at one dose limits one’s ability to detect a potentially more significant dose effect. We, therefore, determined the optimal concentration for increased PTEN expression, by studying our agonists in a dose curve. Cells expressed a basal level of PTEN (0 µM),

which was unchanged when exposed to the EtOH vehicle control (data not shown). When

MCF-7 cells were stimulated with Rosi, we observed a dose-dependent increase in PTEN

expression with a maximal induction occurring at 30 µM (Fig 2.1A; Rosi). When

quantitated to actin levels, we found that Rosi stimulation resulted in a maximum of ~1.5-

fold increase in PTEN levels (Fig 2.1A; Rosi). Similarly, stimulation with Lovastatin

resulted in an increase in PTEN expression (Fig. 2.1B). Induction of PTEN protein levels

began at 1 µM and was maximal at 3 µM, with a ~1-8-fold increase in PTEN (Fig 2.1B:

Lov). This increase in PTEN due to either Rosi or Lovastatin stimulation was also time-

dependent. Stimulation with either Rosi or Lovastatin resulted in an increase in PTEN 26

expression at 24 hr post-stimulation, and maximal effects were observed at 48 hr post stimulation (data not shown). At 72 hrs PTEN levels were near basal. Ciglitazone,

Pioglitazone and Troglitazone did not induce a change in PTEN expression at any time point tested (0-72 hrs; data not shown).

Rosiglitazone induction of PTEN protein expression is PPARγ-dependent

The above data suggest that the specific activation of PPARγ by Rosi, and perhaps Lovastatin, results in an increase in PTEN expression. In order to determine if the effect due to Rosi stimulation was indeed PPARγ-dependent, we utilized the Rosi analog, Compound-66 (Cmpd-66)(57). Analogs to Ciglitazone, Pioglitazone and

Troglitazone were also synthesized, but no analog to Lovastatin was created, due to its complexity. During synthesis of these analogs, a double bond was added to its respective agonist, rendering the compound unable to activate PPARγ, due to steric hindrance.

Analogs to Ciglitazone (∆CG), Pioglitazone (∆PG), and Troglitazone (∆TG) did not induce alterations in PTEN expression in MCF-7 cells, even over a wide range of concentrations and times (data not shown). In contrast to Rosi’s ability to induce PTEN expression, Cmpd-66 was unable to induce PTEN expression to the same extent as Rosi.

Cells stimulated with 3µM of Cmpd-66 had an insignificant ~1.2-fold induction of PTEN

(Fig 2.2; p>0.2). Cmpd-66 is not able to activate PPARγ, therefore these results suggest that Rosi induces PTEN expression in a PPARγ-dependent manner.

Figure 2.1 Lovastatin and Rosiglitazone induce PTEN protein expression in a dose- dependent manner.

Cells were stimulated with Rosiglitazone (A, Rosi) or Lovastatin (B, Lov) as described in

Methods and harvested after 48 hr stimulation as indicated. Levels of PTEN and actin were detected by western blot as described in the Methods. Representative blots of five individual experiments for Rosi stimulation and 3 individual experiments with Lovastatin are displayed. Levels of PTEN were quantitated by densitometry and normalized to actin

(bottom panels). Results are depicted as fold-change compared to nonstimulated cells and are shown in graphical format below the western blots. A) *p<0.001 B) *p<0.001

Figure 2.2 Rosiglitazone’s analog, Compound-66, has a minimal effect on PTEN protein expression.

Cells were stimulated with Cmpd-66 as described in Methods and harvested after 48 hr stimulation as indicated. Levels of PTEN and actin were detected by western blot as described. A) Representative western blots of PTEN and actin (three individual experiments). B) Protein levels were quantitated by densitometry and normalized against actin. Solid bars represent mean PTEN levels (+/- SEM) after stimulation by Rosi (see

Fig. 1 also); Stripped bars represent mean PTEN levels (+/- SEM) after exposure to

Cmpd-66 (*p<0.02) 29

Induction of PTEN protein expression by Lovastatin and Rosiglitazone subsequently inhibits P-AKT and P-p44/42 MAPK levels

The above data indicate that stimulation of MCF-7 cells with Rosi or Lovastatin induces PTEN, but one cannot infer that it is an active protein. Active PTEN is known to inhibit the phosphorylation of AKT, via its lipid phosphatase activity, and phosphorylation of p44/42-MAPK, via its protein phosphatase activity(7). Therefore,

PTEN’s activity is commonly studied by examining the levels of both P-AKT and P- p44/p42. MCF-7 cells have a basal level of P-AKT or P-p44/42 due to normal cellular proliferation (Fig. 2.3A lane 1: control; lane 2: vehicle stimulated). In contrast, stimulation with 30 µM Rosi (Fig. 2.3A; lane 3) or 3 µM Lovastatin (Fig. 2.3A; lane 5) resulted in a decrease in P-AKT, concomitant with PTEN expression, indicating an increase in PTEN lipid phosphatase activity. Furthermore, an inhibition of P-p44/42

MAPK was also observed (Fig. 2.3A; lane 3,5), indicating an increase in PTEN protein phosphatase activity. As expected, stimulation with 3 µM Cmpd-66 (Fig. 2.3A; lane 4) did not alter P-AKT or P-p44/42 MAPK levels. These results are additionally represented in a graphical format in Figure 2.3B. In order to verify that the effect of Lovastatin and

Rosi on P-AKT and P-p44/42 MAPK levels was PTEN-dependent we investigated the ability of Lovastatin and Rosi to induce these changes in BT-549 cells, which are PTEN null. As expected, P-AKT and P-p44/42 MAPK levels did not change in BT549 cells stimulated with either Lovastatin or Rosiglitazone (data not shown). These data suggest that Rosi or Lovastatin stimulation induces functional PTEN protein.

Figure 2.3 Lovastatin and Rosiglitazone inhibit P-AKT and P-MAPK protein expression.

A) Representative blot of P-Akt (top panel), P-p44/p42 (middle panel) and total AKT

(bottom panel) from three individual experiments. MCF-7 cells were either untreated

(lane 1) or treated with EtOH (lane 2), Rosi (lane 3), Cmpd-66 (lane 44) or Lovastatin

(lane 5) for 48 hrs. Cells were harvested and 30 µg of protein extract was subjected to western blot analysis with anti-P-AKT, anti-P-p44/42 or anti-AKT. Both Ponceau S and anti-AKT antibody analysis confirmed equal protein loading. B) Protein levels were quantitated by densitometry and normalized against total AKT. Solid bars represent mean

P-AKT levels after stimulation; Stripped bars represent mean P-p44/42 levels after stimulation. 31

Stimulation of MCF-7 cells with Lovastatin or Rosiglitazone cause G1 cell cycle arrest

Active PTEN has also been shown to result in G1 cell cycle arrest(14). Therefore, in order to confirm PTEN’s activity in the presence of our drugs, we analyzed the cell cycle state of MCF-7 cells after stimulation with either Rosi or Lovastatin. It has previously been shown that sodium butyrate (NaBut) can induce G1 arrest in MCF-7 cells by inhibiting histone deacetylases; therefore NaBut was used as a positive control(38). In order to replicate the environment used in the previous experiments, the cells were not synchronized. As shown in Figure 2.4, unsynchronized cells stimulated with 30 µM Rosi for 48 hrs showed a 10% increase in G1 content (p=0.010). This is comparable to the 15% induction of cells in G1 by NaBut exposure, suggesting that Rosi stimulation results in GI arrest due to increased PTEN expression. In contrast, Cmpd-66 stimulation resulted in a minimal increase of cells in G1. We also observed that

Lovastatin stimulation also resulted in an increase of 13% of cells in G1 (p=0.009). As expected PTEN null cells (BT-549) did not induce cell cycle arrest in response to

Lovastatin or Rosi (data not shown).

Figure 2.4 Lovastatin and Rosiglitazone, but not Compound-66 induce G1 arrest.

MCF-7 cells were either unstimulated (“MCF-7” bar) or stimulated with NaBut, Rosi,

Cmpd-66 or Lovastatin. After a 48-hr treatment, cells were harvested and incubated with propidium iodine as described in the Methods. Cells were then analyzed by flow cytometry. Each bar represents mean % of cells in G1 ± SEM (from three individual experiments). *p=0.009, **p=0.010, NS= not significant

Stimulation with Lovastatin or Rosiglitazone induces PTEN transcription

Our results suggest that stimulation of MCF-7 cells with Rosi increases PTEN in a PPARγ-dependent manner and implies that Lovastatin is also a PPARγ agonist.

However, it is not clear whether this increase in PTEN is due to increased PTEN transcription or decreased protein degradation. We therefore utilized cyclohexamide

(CHX), which inhibits protein synthesis, to determine if either Rosi or Lovastatin stimulation inhibits PTEN degradation. If Rosi or Lovastatin stimulation inhibits PTEN degradation, we would expect to observe no change in PTEN expression when MCF-7 cells, stimulated with Rosi, Cmpd-66 or Lovastatin, are treated with CHX, when compared to Rosi, Cmpd-66, or Lovastatin alone. Co-stimulation with CHX, for 24 or 48 hrs, with either Rosi or Lovastatin, inhibited the production of PTEN, when compared to agonist alone (Figure 2.5A). Cells co-stimulated with CHX and Cmpd-66, for either 24 or

48 hrs, showed little to no change in PTEN production, due to Cmpd-66 only minimally inducing PTEN (Figure 2.5A). We further demonstrated in Figure 2.5A that P-AKT levels decreases due to Rosi stimulation, however its levels do not alter when Rosi and

CHX are used in concert. This suggests that stimulation with Lovastatin and Rosi does not inhibit PTEN protein degradation, but rather it increases its transcription.

Figure 2.5 Lovastatin and Rosiglitazone induce PTEN mRNA expression.

Cells were stimulated and harvested as described in the Methods. A) Cyclohexamide

Treatment. MCF-7 cells were treated with cyclohexamide alone or in concert with either

Rosi or Lovastatin as indicated in the Materials in Methods section. Western Blots analyzed both PTEN and P-AKT levels. Ponceau S analysis confirmed equal protein loading. B) RT-Real time PCR. mRNA levels were quantitated by densitometry and normalized against GAPDH. Unstimulated (MCF-7); EtOH vehicle control; 30 µM Rosi;

3 µM Cmpd-66; 3 µM Lovastatin. Bars represent mean ± SEM of PTEN levels (from

three individual experiments). *p=0.004, **p=0.020

To verify that Lovastatin and Rosi stimulation induces PTEN transcription, we

examined the levels of PTEN mRNA, by RT-PCR, after Rosi or Lovastatin stimulation.

Basal levels of PTEN transcript were observed in unstimulated MCF-7 or cells treated

with EtOH vehicle. In contrast, cells stimulated with 30 µM Rosi for 48 hrs had a ~1.8-

fold increase in PTEN transcript. No induction was detected after stimulation with the

inactive Rosi analog, Cmpd-66, when compared to vehicle control. Furthermore, a ~1.6-

fold induction of PTEN transcript was observed after 48 hr stimulation with 3 µM

Lovastatin (data not shown). Additionally, we investigated the effect of Lovastatin and

Rosi on PTEN message levels by real-time RT-PCR. We found that both Rosi and

Lovastatin induced PTEN mRNA over both EtOH vehicle and Cmpd-66 (Figure 2.5B).

Both Rosi (p=0.004) and Lovastatin (p=0.020) induced ~0.75 fold induction of PTEN,

while Cmpd-66 actually produced a slightly inhibitory affect on PTEN transcript. These

data suggest that Lovastatin and Rosi stimulation results in increased transcription of

PTEN.

Lovastatin stimulation induces PPARγ-specific transcription

Our results indicate that Lovastatin and Rosi induce PTEN transcription.

Additionally, our data with Cmpd-66 (Fig. 2.2) suggest that this effect is mediated by a

PPARγ-dependent mechanism. To date, Rosi has been shown to activate PPARγ- mediated transcription in several models, however Lovastatin has never been shown to be involved in regulating PPARγ. Therefore, in order to definitively demonstrate that

Lovastatin can mediate PPARγ-dependent transcription, we utilized several reporter

assays(84). Rosi was used as a positive control for the following experiments. Luciferase- 36

tagged PPRE was cotransfected into MCF-7 cells with empty vector, wild-type PPARγ,

or dominant-negative PPARγ. These cells were subsequently either unstimulated or stimulated with Rosi, Cmpd-66 or Lovastatin. As expected, Rosi stimulation resulted in a

~900-fold induction of luciferase activity in cells transfected with wild-type PPARγ,

compared to empty vector or non-stimulated cells (Fig. 2.6). In contrast, there was little

activity in stimulated cells transfected with a dominant-negative PPARγ (Fig. 2.6, DN).

Furthermore, stimulation with Cmpd-66 resulted in minimal luciferase activity with all

three vectors, further verifying that this compound is unable to activate PPARγ. In

addition, we found that cells stimulated with Lovastatin had a ~600-fold induction of

luciferase activity when co-transfected with wild-type PPARγ, as compared to empty

vector. Lovastatin stimulation in cells co-transfected with dominant-negative PPARγ

resulted in basal luciferase activity. These results, taken together, confirm that Lovastatin

is signaling through a PPARγ-dependent transcription pathway.

Figure 2.6 Lovastatin and Rosiglitazone induce PPRE regulated transcription.

MCF-7 cells were co-transfected with PPRE-luc and either empty vector (-; solid bars), vector containing wild-type PPARγWT; striped bars) or vector containing dominant- negative PPARγ (DN; spotted bars). Cells were then stimulated with 30 µM Rosi, 3 µM

Cmpd-66 or 3 µM Lovastatin for 48 hrs. The cells were then analyzed for luciferase activity as described. Each bar represents a mean ± SEM of three individual experiments.

*p=0.004, **p=0.013

Lovastatin and Rosiglitazone induce PTEN production in a PPARγ-dependent manner

We further studied the effects of dominant-negative PPARγ on PTEN expression.

Our above data indicate that PPARγ activation due to stimulation with Rosi or Lovastatin induces PTEN in a PPARγ-dependent manner. To concretely verify this, we determined if over-expression of dominant-negative PPARγ would ablate the response observed by

Lovastatin and Rosi stimulation in MCF-7 cells. Figure 2.7A shows that cells transfected with dominant-negative PPARγ and stimulated with Rosi did not induce PTEN, while cells transfected with empty vector or wild-type PPARγ did (~1.5-fold). Additionally, cells transfected with dominant-negative PPARγ did not support Lovastatin-induced

PTEN expression. Cells transfected with wild-type PPARγ or empty vector did, however,

induce PTEN expression (~1.8-fold), indicating that PPARγ is required for Lovastatin-

mediated expression of PTEN. As would be expected, stimulation with Cmpd-66 did not

induce PTEN expression, regardless of transfection.

Our results indicate that both Lovastatin and Rosi induce PTEN expression in a

PPARγ-dependent manner. To further confirm these results, we stimulated mouse

embryonic fibroblasts (MEF) with Rosi, Cmpd-66 or Lovastatin. Two different MEF cell

lines were utilized: a wild-type MEF (MEF-WT) cell line and a PPARγ knock out MEF

cell line (MEF-PPARγ null)(39). Figure 2.7B shows that both MEF-WT and MEF-

PPARγ null cells, which were unstimulated (“MEF”) or stimulated with vehicle control

(“EtOH”) showed basal PTEN expression. We demonstrated that MEF-WT cells

stimulated with Rosi or Lovastatin induced PTEN expression, as similarly observed in

the MCF-7 cell line (p=0.03). However, neither Rosi nor Lovastatin stimulation induced

PTEN expression in the MEF-PPARγ null cell line. In contrast, Cmpd-66 had no effect on PTEN expression in both MEF-WT and MEF-PPARγ null cells (Fig 2.7b). Like PTEN induced by Rosi and Lovastatin stimulation in MCF-7 cells, the PTEN induced in MEF-

WT cells was active as AKT and MAPK phosphorylation decreased concomitant with

PTEN expression (Fig 2.3C). Additionally, cell cycle arrest was observed MEF-WT cells treated with Rosi and Lovastatin compared to unstimulated cells (20.4%, 19.6% respectively; data not shown). Moreover, the MEF-PPARγ null cell line, did not undergo cell cycle arrest or have changes in AKT and MAPK phosphorylation when treated with

Rosi or Lovastatin (data not shown). These data concretely demonstrate that PTEN induction, due to Rosi or Lovastatin, is PPARγ-dependent and not limited to MCF-7 cells.

Figure 2.7 Upregulation of PTEN expression due to Lovastatin and Rosiglitazone is

PPARγ-dependent and not cell line specific.

A) MCF-7 cells were transfected with empty vector (-; solid bars), vector containing

wild-type PPARγ (WT; striped bars) or vector containing dominant-negative PPARγ

(DN; spotted bars). *p<0.001, **p=0.001. B) MEF-WT (solid bars) and MEF-PPARγ

null (striped bars) cell lines were also analyzed. The MEF-WT and MEF-PPARγ null cell

lines were either not stimulated (1) or stimulated with EtOH (2), 30 µM Rosi (3), 3 µM

Cmpd-66 (4), or 3 µM Lovastatin (5) for 48 hrs. Protein content was analyzed for PTEN

expression by western blot (expressed as mean +/- SEM from 3 individual experiments)

as described in the Methods section. Results are depicted in a quantitated graphical

format. *p=0.03 C) The MEF-WT cell line was either not stimulated (1) or stimulated

with EtOH (2), 30 µM Rosi (3), 3 µM Cmpd-66 (4), or 3 µM Lovastatin (5) for 48 hrs.

Protein content was analyzed for both P-AKT and P-p44/42 expression by western blot

(expressed as mean +/- SEM from 3 individual experiments).

2.4 Discussion

Our current observations demonstrate that stimulation of MCF-7 cells with 30 µM

Rosi or 3 µM Lovastatin induces PPARγ-mediated transcription of PTEN, subsequently increasing PTEN protein levels. Furthermore, we show that stimulation with either Rosi or Lovastatin, induces a PTEN protein that is both protein- and lipid-phosphatase active.

We also demonstrate that the induction of PTEN due to Rosi or Lovastatin is a PPARγ- dependent mechanism through the use of Cmpd-66, dominant negative PPARγ and

PPARγ-null MEF’s. Perhaps more importantly, we have demonstrated for the first time, that Lovastatin increases PTEN via PPARγ-mediated transcription, thus suggesting that

Lovastatin may also function as an anti-proliferative agent(40). Taken together, these data provide further evidence that PPARγ is a tumor suppressor and that one of its tumor suppressing mechanisms is upregulating the transcription of PTEN.

In 2001, Patel and co-workers’ report suggested that Rosi induces PTEN expression in MCF-7 cells, it however remained correlative, as the mechanism of induction was not investigated fully. We demonstrate here, using biochemical and molecular techniques, that stimulation of MCF-7 and MEF-WT cells with either

Lovastatin or Rosi induces PPARγ-mediated transcription of PTEN. Previous work has been in macrophages leaving the question open if what is observed in one cell line would equally occur within another. It is now well known that TZD’s have cell line-dependent functions, and indeed, we demonstrate that phenomenon here with the other TZD’s tested. We have previously shown that increases in PTEN protein does not necessarily

correlate with increases in down-stream PTEN-mediated cellular events(89, 90), thus, it is imperative to determine if the increased PTEN protein is active. Patel and co-workers did not analyze the normal readouts of PTEN’s activity such as the phosphorylation state of AKT and MAPK, or G1 arrest. Additionally, they did not determine if other TZD’s have a similar affect in MCF-7 cells. Lastly, and of most significance, it has not been demonstrated, to date, that PTEN induction by Rosi is dependent upon the presence of

PPARγ.

Our data indicate that while the TZD PPARγ agonist Rosi has the ability to induce

PTEN in MCF-7 cells, the other TZD’s Ciglitazone, Pioglitazone and Troglitazone do not. Until recently, it has been thought that all the TZD’s function in a similar manner.

However, it is now becoming more apparent that each agonist may have its own cellular functions, some of which may be PPARγ-independent(41). Our current data with Rosi support this idea. Additionally, pharmacological affinity studies have previously shown that of the 4 TZD’s; Rosi has the highest affinity for PPARγ, followed by Pioglitazone>

Troglitazone> Ciglitazone(57, 58).

Previously, PPARγ agonists, in particular Rosi, have been shown to correlate with

PTEN expression in MCF-7 breast cancer cells(21), macrophages(21), monocytes(92), bronchoalveolar lavage cells(42) and AsPC-1 pancreatic cancer cells(43). We show here that stimulation of MCF-7 cells with 30 µM Rosi results in a ~1.5-fold increase in PTEN.

This level of induction is in agreement with the ability of other transcription factors to induce PTEN expression. Egr-1 activation has been shown to induce PTEN at a ~1.6-fold level(44). In addition, the level of induction that we observe with Rosi is comparable to

PTEN induction by Rosi in monocytes, where Rosi stimulation results in a ~1.4-fold 44

increase in PTEN. Moreover, a modest increase in PTEN expression was also shown in both the bronchoalveolar lavage and AsPC-1 cell models. Interestingly, Rosi stimulation in macrophages results in a ~10-fold increase in PTEN(21). This may suggest that macrophages respond to Rosi stimulation in a PPARγ-independent manner, which does not occur in other lines, or that PPARγ signaling is higher in macrophages compared to other cell lines. Nevertheless, our data strongly demonstrate that Rosi stimulates PTEN expression in MCF-7 cells in a PPARγ-dependent manner. Furthermore, our data also indicate that Rosi’s ability to induce PTEN expression is not cell line dependent.

However, in order to confirm these data, we tested Rosi’s ability to induce PTEN in three different cell lines: MEF-WT, MEF-PPARγ null and BT-549. Our results demonstrate that both Lovastatin and Rosi can induce PTEN expression in MCF-7 cells and the MEF-

WT cell line. However, while PTEN was induced in MCF-7 and MEF-WT’s, there was no induction of PTEN protein in response to Lovastatin or Rosi in the MEF-PPARγ null cell line. This indicates that PPARγ is necessary for the induction of PTEN due to Rosi or

Lovastatin stimulation and that this induction of PTEN is not cell line specific. We have expanded upon the previously published MCF-7 work(21) and demonstrate here that 30

µM Rosi produced the maximal induction of PTEN expression in MCF-7 cells. These results are in concordance with the ability of Rosi to stimulate BRCA1 expression in

MCF-7 cells(45). Both PTEN (shown here) and BRCA1 expression is maximal at 30 µM

Rosi. Like Patel and co-workers(21), we see an increase in PTEN expression at 1 µM

Rosi; however, we found that 30 µM Rosi provides the maximal effect. Additionally,

Patel and co-workers did not examine other concentrations in the MCF-7 cell line.

Furthermore, in the BT-549 PTEN null breast cancer cell line, we did not observe an

increase in PTEN expression or an inhibition of P-AKT and P-MAPK after Rosi or

Lovastatin stimulation. This indicates that the response we observed in MCF-7 cells is

due to the production of PTEN and not a non-specific response to Rosi or Lovastatin. In

contrast, we were able to observe the inhibition of cyclin D1 in the BT-549 cell line with

Rosi or Lovastatin stimulation (data not shown) suggesting that the cyclin D1 response to

Lovastatin and Rosi in not dependent upon an increase in PTEN protein expression.

The TZD’s are used clinically due to their high affinity for PPARγ, but recent data suggest they may play secondary PPARγ-independent roles(41). However, to date, it has been assumed that Rosi is signaling in a PPARγ-dependent manner to induce PTEN, therefore we utilized Cmpd-66 to determine PPARγ-specificity. Previously Cmpd-66 has been shown to only minimally compete with Rosi and have little affinity for PPARγ, demonstrating that it functions in a PPARγ-independent manner(57). Due to Cmpd-66’s ability to minimally compete with Rosi, a small induction of PTEN with Cmpd-66 stimulation should not be unexpected. Our analog data was then further confirmed through the use of PPARγ plasmids demonstrating that both Lovastatin and Rosi can induce PTEN in a PPARγ-dependent manner.

It is well known that PTEN can be maintained at an inactive state within the cell for an extended period to time (44); therefore it is important to determine if Rosi-induced

PTEN is active. We were able to demonstrate an increase in PTEN phosphatase activity after Rosi stimulation. This decrease in P-AKT and P-MAPK levels within the cell thereby stops cellular proliferation, induces apoptosis and causes G1 arrest. Our data

therefore indicate that the inhibition of cellular proliferation in MCF-7 cells after

stimulation with Rosi, as seen in previously published data(21), is due to an increase in

active PTEN.

Lovastatin-simulated MCF-7 cells resulted in decreased P-AKT and P-MAPK

levels, and increased the percentage of cells in G1. These data resemble previously

published data, where Lovastatin was shown to increase the percentage of cells in G1

using the MCF-7 model(46). These data suggest that the increased G1 arrest after

Lovastatin stimulation could also be due to the increase in active PTEN.

Until recently, the focus on Lovastatin has been on its ability to regulate

cholesterol levels. However, in leukemia and breast cancer cells, Lovastatin was

implicated as a regulator of the MAPK pathway and the cell cycle(98, 99), respectively

Other statins, including atrovastatin and fluvastatin, have pro-apoptotic functions within

MCF-7 cells, suggesting that the statins, as a group, possess properties that may be

beneficial in the treatment of breast cancer patients(47). Interestingly, women taking

statins have a 68% reduction in the risk of breast cancer(40). Work in Sprague-Dawley

rats has demonstrated that atorvastatin treatment increases PTEN protein expression(48).

This suggests that the statins as a group may induce PTEN expression. We now gather

the first evidence that a potential mechanism for cellular regulation by Lovastatin is

through PPARγ. Our data indicates that Lovastatin stimulation results in the activation of

PPARγ-mediated transcription. Currently, it is unclear, in the Sprague-Dawley rat model, if the role of atrovastatin is through PPARγ mediated transcription. However, in light of

our data, it is possibile that all or most statins induce PPARγ, and subsequently PTEN,

opening up an exciting avenue of investigation for both cardiac and cancer research. 47

The mechanism behind PTEN induction, due to PPARγ agonists, has yet to be fully understood. Two molecular connections between PTEN and PPARγ have been previously hypothesized: PPARγ regulates the transcription of PTEN(21) and/or PPARγ regulates a secondary factor, which regulates PTEN(102). Furthermore, because PTEN levels can be regulated by both an increase in transcription or a decrease in degradation(44, 89), it remained a possibility that PPARγ agonists induced PTEN expression by decreasing protein degradation. Our data indicate that both Lovastatin and

Rosi stimulation do not inhibit PTEN degradation, but rather induce PTEN transcription.

The specific regions in the promoter that is required for PPARγ mediated transcription by

Lovastatin and Rosi remain to be elucidated and are beyond the scope of this paper.

While two putative PPRE’s have been suggested(21) these sites are well upstream of the minimal PTEN promoter and it remains unclear what role, if any, these sites have on

PTEN transcription. We have been unable to demonstrate that these are indeed Lovastatin and Rosi responsive PPARγ binding sites in breast cancer cell lines (data not shown).

Nonetheless PTEN’s minimal promoter has several sites that are potential candidates for

PPARγ mediated transcription mediated by Lovastatin and Rosi. One must also keep in mind that these data do not limit PPARγ’s ability to regulate PTEN only through inducing PTEN transcription and it remains a possibility that PPARγ is mediating the transcription of another transcription factor that binds directly to the PTEN promoter, this remains to be determined.

In conclusion, we have demonstrated that PPARγ directly regulates PTEN at the transcriptional level. Furthermore, since PTEN is constitutively active, it may be

worthwhile to examine the effects of Lovastatin and Rosi as a mechanism for increasing

PTEN expression for both the treatment and prevention of cancer. Indeed, the data with

the Sprague-Dawley rats may begin to suggest that treatment of patients (eg, CS) or

cancers that have hemizygous deletions or haploinsufficiency of PTEN with Lovastatin or

Rosi may be conceivable. Despite these encouraging results, we must be aware that these medications may indeed harm patients with germline intragenic PTEN mutations or those with neoplasias with somatic intragenic mutations by raising levels of mutant protein.

CHAPTER 3

REGULATION OF THE PTEN PROMOTER BY STATINS AND SREBP

3.1 Introduction

Germline mutations in PTEN, a tumor suppressor gene on 10q23, occur in 85% of patients with the autosomal dominant CS [MIM 158350](14, 17, 64). This syndrome is reported to affect approximately 1 in 200,000 individuals. However, because CS is difficult to diagnose, this is generally thought to be an underestimate(30). Patients diagnosed with CS have a 25-50% lifetime risk of developing female breast cancer compared to ~13% in the general population(29, 49). In addition to breast cancer, CS patients also have thyroid neoplasias, mucocutaneous lesions, fibrocystic breast disease, uterine leiomyoma and macrocephaly(2, 67). Germline PTEN mutations are also associated with subsets of patients with developmental disorders. Approximately 65% of patients with BRRS [MIM 153480], 20% of those with Proteus syndrome [MIM 176920],

50% of a Proteus-like syndrome(30) and 10-20% of autism spectrum disorder with macrocephaly(14, 16) share germline PTEN mutations as an etiology and have all been classified as PHTS(5). Furthermore, somatic alterations in PTEN, whether by genetic or epigenetic mechanisms, play some role in the pathogenesis of a broad range of solid tumors, such as sporadic carcinomas of the breast, thyroid, endometrium and colon(30). 50

PTEN’s protein product, PTEN, is a dual-specificity phosphatase with both lipid

and protein phosphatase activities(31), which elicits cell cycle arrest and apoptosis.

PTEN is a constitutively expressed protein, therefore, regulation of its protein levels

through transcription is key to its function. In recent years, transcriptional regulation of

PTEN has been researched more extensively, however, there is still much to be

understood. Several groups have shown that PTEN transcription may be regulated by

Egr-1(51), p53(20), Sp1(33), NF-κB(26), CBF-1(24), USF1(23) and c-Jun(25).

Additionally, in 2001, two putative binding sites for the transcription factor PPARγ were identified approximately 10kb upstream of the minimal promoter region of PTEN (21); specific binding of PPARγ was later confirmed(50). Moreover, our laboratory demonstrated that PPARγ, through activation by Rosiglitazone or Lovastatin, induces

PTEN transcription and subsequently upregulates PTEN protein levels(51). These data were the first to suggest that a statin, more specifically Lovastatin, signals through

PPARγ and upregulates PTEN expression.

Statins have long been thought to have some anti-carcinogenic properties, but concrete evidence remains to be lacking. Clinically, statins are used as HMG-CoA inhibitors, which subsequently downregulate cholesterol production through the mevalonate pathway. However, further research shows that this pathway can also regulate other proteins, such as Ras and Rho, which are key proteins involved in cancer development(27). Lovastatin (Mevacor), Simvastatin (Zocor), Pravastatin (Livals), and

Fluvastatin (Lescol) are commonly prescribed for millions of patients to aid in lowering cholesterol levels for both the primary and secondary prevention of cardiovascular disease(28). In this study, we investigated if four statins (Simvastatin, Pravastatin, 51

Fluvastatin, and Mevastatin), beyond Lovastatin, have the ability to act as anti-

carcinogenic agents by upregulating PTEN expression through PPARγ, similar to

Lovastatin or via other mechanisms.

3.2 Materials and Methods

Materials

Fluvastatin, Mevastatin, Lovastatin, Pravastatin and Simvastatin were obtained

from Cayman Chemical (Ann Arbor, MI). ALLN was purchased from Biomol

International (Plymouth Meeting, PA). Antibodies were obtained from Cascade

Bioscience, Waltham, MA (PTEN 6H2.1), Sigma-Aldrich (Actin), and Cell Signaling

(PPARγ and SREBP1). Cell culture media was obtained from the Cleveland Clinic Media

Core. M-PER Mammalian Protein Extraction Reagent was obtained from Pierce

Biotechnology Inc. (Rockford, IL). All other reagents were purchased from standard

commercial sources.

Cell culture and stimulation

The MCF-7 cell line was maintained at 37°C with 5% CO2 in DMEM containing

10% FBS and 100 units/ml each Pen/Strep. MDA-MB-231, MDA-MD-435 and T47D

breast cancer cell lines were maintained at 37°C with 5% CO2 in RPMI containing 10%

FBS and 100 units/ml each Pen/Strep. Cells were plated at 1.0 x 108 24 hrs prior to treatment. After 24 hrs, cells were treated as indicated in results section and figure legends. Cells were subsequently incubated for an additional 48 hrs prior to harvesting. 52

Protein Extraction

MCF-7, MDA-MB-231, MDA-MB-435 and T47D cells were plated and stimulated as described above. At the time of harvest, media was removed and the cells were washed with PBS. Cells were then harvested into M-PER lysis buffer containing

PMSF (0.75mg/ml), benzamidine hydrochloride (0.5 mg/ml), leupeptin (2µg/ml), aprotinin (2 µg/ml), pepstatin (2 µg/ml), β-glycerophosphate (10 mM), NaOV (0.2 mM), and NaF (25mM). Cells were incubated at room temperature with lysis buffer for one minute before harvesting by scraping. Samples were then centrifuged at 16,000 X g for

10 min at 4ºC to remove cellular debris. The resulting supernatant was stored at -80°C.

Protein concentration was determined using the bicotinic method(35) using BSA as a standard.

Western Blot

Proteins (30µg) were prepared by the Laemelli method(36), then separated on a

10% SDS-PAGE gel, and electrophoretically transferred onto nitrocellulose. Equal protein loading between conditions was confirmed by staining with Ponceau S solution.

Non-specific binding was blocked by incubating the nitrocellulose blots with 5% milk in

TBS-T (100mM Tris, pH, 1M NaCL, 1% Tween-20) for 1 hr at room temperature. Blots were then incubated with the primary antibody (1:1000 in 3% BSA) for 2 hrs at room temperature. Following the primary incubation, the blots were washed with TBS-T for 1 hr with frequent changes of buffer. Blots were then incubated with the appropriate secondary antibody conjugated to horseradish peroxidase (Promega; Madison, WI)

(1:2500 dilution in 5% milk) overnight at 4°C, and washed with TBS-T for 1 hr. Protein 53

bands were visualized using enhanced chemiluminescence as described by the manufacturer (Amersham Pharmacia Corp; Piscataway, NJ). The resultant films were then quantified using NIH-Imager densitometry software.

RT-PCR

MCF-7 cells were stimulated as described above. After stimulation, cells were released by trypsin treatment and subsequently washed three times with PBS through centrifugation. Total RNA was extracted from cells following the Gentra Versagene RNA

Purification System Protocol (Minneapolis, MN) and then converted to cDNA by

Superscript II Reverse Transcriptase after DNase treatment. The resultant cDNA was subjected to multiplex PCR amplification using primers specific to PTEN exon 3 and exon 5 (F: 5’ TGGATTCAAAGCATAAAAACCA 3’; R: 5’

AAAAGGATATTGTGCAACTCTGC 3’) and β-actin (Quantum RNA β-actin; Ambion

INC; Austin, TX). Primers were allowed to anneal at 55°C for 28 cycles. The products from the PCR reactions were run on a 1% agarose gel containing ethidum bromide and visualized under a UV light.

Reporter Assay

Plasmids were co-transfected into 6-well cultures of MCF-7 cells with 3µl/well of

FuGene (Roche). Each well was co-transfected with 500 ng of the reporter plasmid

PPRETKLUC, 50 ng Renilla luciferase control plasmid and 100 ng of receptor expression vector (pcDNA3, WT PPARγ1, or L468A/E471A PPARγ1). Each plasmid has previously been described(84, 104). After 12 hrs, cells were stimulated for 48 hrs. 54

PTEN Promoter was PCR amplified from normal genomic DNA and subsequently

cloned into a TOPO-TA vector. DNA was PCR amplified using 55°C as the annealing

temperature for 30 cycles. All PCR amplification products were verified by direct DNA

sequencing (ABI 3730xl DNA Analyzer) and positive clones were subcloned into a

pGL3.1-Basic vector (Promega, Madison, WI). To determine luciferase activity, 6-well

plates were co-transfected with 1 µg/well of a pGL3-PTEN construct, and 50 ng/well

Renilla luciferase control plasmid with 3µl/well of FuGene. After 12 hrs, cells were

stimulated with EtOH, 3 µM Lovastatin, 3 µM Simvastatin, 30 µM Pravastatin or 18 µM

Fluvastatin for 48 hrs. Cells were harvested with 1X passive luciferase lysis buffer (Dual-

Luciferase Reporter Assay System; Promega) and analyzed on a luminometer (LMax

11384; Molecular Devices) using Renilla luciferase as an internal transfection control.

Electromobility Shift Assay (EMSA)

PTEN promoter sequence was isolated through PCR amplification from normal genomic

DNA. The DNA was PCR amplified with 30 cycles at the annealing temperature of 55°C in 20 µl reactions using HotStar and Q Solution (Qiagen). Each of the above products was radiolabeled with 32P-γATP via T4 kinase. To examine DNA-protein interaction, 1

ng of radiolabeled probe was incubated with 2 µg of either untreated or statin-treated

nuclear protein extract for 20 min at room temperature with binding buffer containing 10

mM HEPES (pH 7.5), 2.5 mM MgCl2, 50 mM NaCl, 0.5 mM DTT, 4% glycerol, 1

µg/mL BSA, and 2 µg poly dI/dC. Unlabeled probe in 5X molar excess was used as the

specific competitor, while a random oligonucleotide sequence was used as the

nonspecific competitor. DNA-protein complexes were resolved on a 4% nondenaturing

PAGE gel at 150 V for 3.5 hrs at 4°C and visualized using a Phospho-Imager (Amersham

Biosciences, Piscataway, NJ).

Statistical Analysis

Statistical analysis was done using Student’s t test. Data are mean ± SD of three independent experiments and normalized to a control. P<0.05 is considered significant.

3.3 Results

Simvastatin, Pravastatin, and Fluvastatin induce PTEN protein expression in a dose- dependent manner

We have recently demonstrated that Lovastatin upregulates PTEN expression(51).

To determine if other statins could upregulate PTEN expression, we stimulated MCF-7 breast cancer cells with four additional statins: Simvastatin, Pravastatin, Fluvastatin, and

Mevastatin. The optimal concentration for increased PTEN expression was determined by performing a dose curve based on their previously established IC50’s in breast cancer cell lines(52). MCF-7 cells expressed a basal level of PTEN, which was unchanged when exposed to the EtOH vehicle control (Fig. 3.1; 0 µM). When MCF-7 cells were stimulated with Mevastatin, we did not observe any changes in PTEN expression, despite the wide range of doses centered on its IC50 (Fig. 3.1D; 0-25 µM). In contrast,

Simvastatin, Pravastatin, and Fluvastatin induced a dose-dependent increase in PTEN expression after treatment. The greatest increase in PTEN expression occurred at 3 µM, 56

30 µM, and 18 µM for Simvastatin, Pravastatin and Fluvastatin, respectively (Fig. 3.1A-

C). Our previous work demonstrated that 3 µM Lovastatin could stimulate ~1.8-fold induction of PTEN expression. Our current study exhibits a similar induction of PTEN with Simvastatin (~1.5-fold), Pravastatin (~1.4-fold) and Fluvastatin (~1.6-fold).

Statin treatment induces PTEN mRNA expression

Our previous work indicated that Lovastatin induces PTEN protein by upregulating PTEN transcription, and subsequently, its mRNA levels(51). To determine if Simvastatin, Pravastatin, Fluvastatin and Mevastatin stimulation induces PTEN

transcription in a similar manner as Lovastatin, we examined the levels of PTEN mRNA

by RT-PCR after treatment. Basal levels of PTEN transcript were observed in

unstimulated MCF-7 cells or those treated with EtOH vehicle (Fig. 3.2A; EtOH). In

contrast, cells stimulated with 3 µM Simvastatin for 48 hrs had a ~1.7-fold increase in

PTEN transcript (Fig. 3.2A; Sim). An ~1.8-fold induction of PTEN transcript was

observed after 30 µM Pravastatin stimulation (Fig. 3.2A; Pra). Furthermore, an ~1.8-fold

induction of PTEN transcript was also observed after stimulation with 18 µM Fluvastatin

(Fig. 3.2A; Flu). The induction was observed with these statins are very similar to that

observed with Lovastatin (~1.8-fold)(51). We additionally examined PTEN mRNA levels

in MCF-7 cells after stimulation with Mevastatin to confirm its inability to induce PTEN

expression. As expected we did not observe an induction of PTEN mRNA after

Mevastatin treatment (Fig. 3.2A; Mev). These data demonstrate that Simvastatin,

Pravastatin, and Fluvastatin stimulation induces PTEN transcription.

Figure 3.1 Simvastatin, Pravastatin and Fluvastatin induce PTEN protein expression in a dose-dependent manner.

Cells were stimulated with Simvastatin A), Pravastatin B) Fluvastatin C), or Mevastatin

D) and harvested after 48 hr stimulation as indicated. Levels of PTEN and actin were detected by western blot as described in the Methods. Representative blots of three individual experiments are displayed.

Figure 3.2 Simvastatin, Pravastatin and Fluvastatin induce PTEN mRNA expression and

PPRE regulated transcription.

Cells were stimulated and mRNA harvested as described in the Methods. A) PTEN mRNA levels were quantitated by densitometry and normalized against actin [EtOH vehicle control; 3 µM Simvastatin (Sim); 30 µM Pravastatin (Pra); and Fluvastatin (Flu)].

Bars represent mean ± SEM of PTEN levels (from three individual experiments). B)

MCF-7 cells were co-transfected with PPRE-Luc and either empty vector (-; gray bars),

wild-type PPARγ (WT; black bars) or dominant-negative PPARγ (DN; white bars). Cells

were then stimulated with 3 µM Lovastatin, 3 µM Simvastatin, 30 µM Pravastatin, 18

µM Fluvastatin or 18 µM Mevastatin for 48 hrs and analyzed for luciferase activity as

described. Each bar represents a mean ± SEM of three individual experiments.

Statin stimulation induces PPARγ-mediated transcription

We previously demonstrated that Lovastatin induces PTEN and upregulates

PPARγ-mediated transcription similar to Rosiglitazone, a known synthetic PPARγ ligand(51). To determine whether Simvastatin, Pravastatin, and Fluvastatin could regulate

PPARγ-mediated transcription as well, we utilized a previously described reporter assay system(84, 104). This reporter assay system takes advantage of the known PPARγ response element (PPRE), which has been luciferase-tagged(53). MCF-7 cells were co- transfected with a PPRE construct and empty vector, WT-PPARγ, or DN-PPARγ. Twelve hrs later, these cells were treated with EtOH, Lovastatin, Simvastatin, Pravastatin,

Fluvastatin, or Mevastatin for 48 hrs. As expected, Lovastatin stimulation induced

PPARγ-mediated transcription ~100-fold over EtOH-treated MCF-7 cells (Fig. 3.2B;

Lov-WT). Simvastatin treatment resulted in an ~80-fold induction of luciferase activity in cells transfected with WT-PPARγ, compared to EtOH-stimulated cells (Fig. 3.2B; Sim-

WT). Treatment with Pravastatin resulted in activation of PPARγ-mediated transcription with a ~70-fold induction (Fig. 3.2B; Pra-WT). In addition, we observed that cells stimulated with Fluvastatin had a ~120-fold induction of luciferase activity when co- transfected with WT-PPARγ, as compared to EtOH-treated cells (Fig. 3.2B; Flu-WT).

Unexpectedly, Mevastatin stimulation induced PPARγ-mediated transcription ~70-fold over EtOH-treated cells (Fig. 3.2B; Mev-WT). In contrast, there was little activity in cells treated with any of the statins after transfection with a DN-PPARγ (Fig. 3.2B; DN). The above results indicate that all five statins can both significantly and specifically induce

PPARγ-mediated transcription. These data were both unexpected and interesting. We

observed that Lovastatin, Simvastatin, Pravastatin, and Fluvastatin induced PTEN expression, while Mevastatin did not (Fig. 3.1). However, the above reporter assay results indicate that all five statins can signal though PPARγ-mediated transcription. These results were unexpected because our previously published data demonstrated that PPARγ protein is necessary to induce PTEN(51).

We initially expected to observe all the statins induce PTEN expression signaling through PPARγ-mediated transcription, however, our Mevastatin results suggest that this may not be the signaling pathway utilized. It has been previously suggested that statins can signal through sterol response element binding protein (SREBP), however, the exact mechanisms are still being elucidated(27, 54). Additionally, published data indicate that

SREBP can upregulate PPARγ protein expression and subsequently induce PPARγ- mediated transcription(55). Therefore, based on our above results and these published data, we hypothesized three potential model pathways of PTEN upregulation by statins via SREBP and PPARγ (Fig. 3.3). In Model A, statins signal through SREBP to induce

PPARγ-mediated transcription, subsequently increasing PTEN expression (Fig. 3.3A).

However, it is entirely possible that statins may upregulate PPARγ-mediated transcription, and therefore, PTEN through a signaling pathway distinct from that of

SREBP (Fig. 3.3B, Model B). Our results from the above reporter assay may indicate that statins may induce PTEN expression through a signaling pathway that utilizes PPARγ activity independent of PPARγ’s transcriptional activity (Fig. 3.3C, Model C).

Figure 3.3 Three potential models of statin-mediated PTEN upregulation.

A) Statins regulate SREBP levels, which induce PPARγ-mediated transcription. PTEN transcription is subsequently increased. B) Statins and SREBP can both induce PPARγ- mediated transcription, however, their mechanisms are independent of each other. PTEN transcription is subsequently increased. C) Statins regulate PPARγ protein activity, which regulates a transcription factor that induces PTEN transcription. SREBP also induces

PTEN transcription, but through regulation of PPARγ-mediated transcription.

Both statins and SREBP can induce PTEN protein expression

As noted above, published data suggest that statins upregulate PTEN through

PPARγ-mediated transcription, and that SREBP may be a mediator (Fig. 3.3A)(55). To test this hypothesis, we utilized N-acetyl-leucyl-leucyl-norleucinal (ALLN), which inhibits SREBP catabolism, thus upregulating its expression and function. MCF-7 cells were treated with Lovastatin, Simvastatin, Pravastatin, Fluvastatin, or ALLN for 48 hrs and whole cell protein lysates were examined by western blot analysis. Vehicle (EtOH)- treated cells express a basal level of SREBP and PPARγ protein (Fig. 3.4A, lane 1

labelled -). Cells stimulated with Lovastatin (L), Simvastatin (S), Pravastatin (P), and

Fluvastatin (F) do not significantly alter SREBP or PPARγ protein expression (Fig. 3.4A,

lanes 2-5). In contrast, ALLN treated cells (A) exhibit an increase in both SREBP and

PPARγ protein expression as expected (Fig. 3.4A, lane 6).

Our current study indicates that statins can universally induce PTEN expression and suggest that SREBP can as well. Therefore, to establish that the effects we see with statin stimulation are not cell-specific, and that SREBP can induce PTEN expression, we compared our results obtained in MCF-7 (Fig. 3.2B, top panel) with three other breast cancer cell lines: MDA-MB-435 (second panel), MDA-MB-231 (third panel) and T47D

(fourth panel). While the basal level of PTEN expression varies across the cell lines, statin and ALLN stimulation results in ~1.8-fold increased PTEN expression in all four cell lines (Fig. 3.4C).

Figure 3.4 PTEN upregulation by statins and ALLN are not cell line-specific.

A) Cells were stimulated with EtOH (-), Lovastatin (L), Simvastatin (Sim), Pravastatin

(Pra) or Fluvastatin (Flu) in the presence or absence of ALLN (A) as described in

Methods. Protein was harvested after 48 hr stimulation and levels of SREBP, PPARγ and actin were detected by western blot. Representative blots of three individual experiments are displayed. B) MCF-7, MDA-MB-435, MDA-MB-231 and T47D breast cancer cell lines were stimulated with 3 µM Lovastatin (L), 3 µM Simvastatin (S), 30 µM

Pravastatin (P), 18 µM Fluvastatin (F), or 15 µM ALLN (A) for 48 hrs. Levels of PTEN were detected by western blot as described in the Methods and Ponceus S indicated even

loading. Representative blots of three individual experiments are displayed. C)

Quantification of PTEN protein normalized to actin levels. Western blot results are depicted as fold-change and are shown in graphical format: EtOH, Lovastatin (Lov),

Simvastatin (Sim), Pravastatin (Pra), Fluvastatin (Flu) and ALLN. Each bar represents a mean ± SEM of three individual experiments

SREBP induction of PPARγ-mediated transcription antagonized by statins

ALLN upregulation of SREBP and PPARγ expression is thought to result in an increase in PPARγ-mediated transcription(55). To determine if upregulation of SREBP can indeed induce PPARγ-mediated transcription in our system, we performed a PPARγ reporter assay with cells treated with ALLN. In addition we observed the effect of costimulation of statin/ALLN (Lov/ALLN, Sim/ALLN, Pra/ALLN, Flu/ALLN) to determine if the agonists have an additive affect. ALLN stimulation of MCF-7 cells in the presence of WT-PPARγ significantly induced PPARγ-mediated transcription (~160-fold;

Fig. 3.5A, ALLN-WT), which was inhibited when treated in the presence of DN-PPARγ

(~20-fold; Fig. 3.5A, ALLN-DN). Interestingly, when MCF-7 cells were treated with

ALLN in concert with the any of four statins that induced PTEN expression (Lovastatin,

Simvastatin, Pravastatin, and Fluvastatin), we observed an inhibition of luciferase activity compared to ALLN alone. By comparing these results (Fig. 3.5A) with our previous reporter assay results (Fig. 3.2B), we observed a significant inhibition of PPARγ- mediated transcription when MCF-7 cells are treated with a statin/ALLN combination compared to statin treatment alone. Co-stimulation of Lovastatin and ALLN (Lov/ALLN)

inhibited ALLN’s induction by 50% and Lovastatin’s induction by 20%, while treating

cells with Simvastatin and ALLN (Sim/ALLN) inhibited their induction by 40% and

70%, respectively. Treatment of Pravastatin and ALLN (Pra/ALLN) in concert inhibited

their induction by 15% and 60%, respectively. Additionally co-stimulation of Fluvastatin

and ALLN (Flu/ALLN) inhibited Fluvastatin’s induction 70% and ALLN’s induction

75%.

These observations suggest that ALLN-related upregulation of PPARγ-mediated

transcription may be due to SREBP induction, however, it may also be a result of

ALLN’s proteasome inhibitor activity. Therefore, to test ALLN’s role in PPARγ- mediated transcription, we performed a reporter assay (similar to that shown in Fig. 3.5A) with MG-132, another proteasome inhibitor(17). In contrast to ALLN, MG-132 did not produce an induction of PPARγ-mediated transcription indicating that this is not a

general proteasome inhibitor phenomenon (data not shown). These data suggest that

SREBP induces PPARγ protein, resulting in increased PPARγ transcriptional activity.

Additionally, these data provide evidence that statins do not signal through SREBP to

induce PTEN expression, thus, excluding Model A (Fig. 3.3A). Instead, our data suggest

that these two signaling pathways appear to antagonize each other.

Figure 3.5 SREBP induces PPARγ-mediated transcription, which is inhibited by the

statins.

A) MCF-7 cells were transfected with PPRE-Luc. After 12 hrs, cells were stimulated

with 15 µM ALLN alone or in combination with 3 µM Lovastatin (Lov), 3 µM

Simvastatin (Sim), 30 µM Pravastatin (Pra), or 18 µM Fluvastatin (Flu) for 48 hrs and

analyzed for luciferase activity as described. Each bar represents a mean ± SEM of three

individual experiments. B) MCF-7 cells were transfected with PPRE-Luc and stimulated

with 3 µM Simvastatin (Sim), 30 µM Pravastatin (Pra), 18 µM Fluvastatin (Flu), 15 µM

ALLN or a combination of the statins with ALLN for 48 hrs. Cells were then analyzed

for luciferase activity as described. Each bar represents a mean ± SEM of three individual

experiments. 67

SREBP, but not the statins, induce PPARγ-mediated transcription to increase PTEN

protein expression

Our above results suggest that statins signal down a pathway separate from

SREBP, potentially as illustrated in Model B (Fig. 3.3B). Additionally, SREBP may have

the ability to induce PTEN expression through the upregulation of PPARγ transcriptional

activity although this observation may be artificial. In the above reporter assay system

(Fig. 3.2B and Fig. 3.5A), we added exogenous PPARγ protein to observe an increase in

PPARγ-mediated transcription. However, in our initial western blots (Fig. 3.1), we observed an induction of PTEN expression after statin stimulation without the addition of exogenous PPARγ. In order to determine if the statins and SREBP truly signal through

PPARγ to induce PTEN transcription, we performed a reporter assay in the absence of exogenous PPARγ protein. MCF-7 cells were transfected with only the PPRE-Luc vector and subsequently stimulated with the four statins, ALLN or the statin/ALLN combination. Statin stimulation alone presented no induction of PPARγ-mediated transcription, compared to EtOH, when exogenous PPARγ was not added to the system

(Fig. 3.5B; bars 1-5). In contrast, we observed an induction of luciferase activity after

ALLN stimulation (Fig. 3.5B; bar 6; ~40-fold). Stimulation of MCF-7 cells with ALLN

in concert with the statins altered PPARγ-mediated transcription compared to ALLN

alone, but remained significantly induced over statin treatment alone (Fig. 3.5B; bars 7-

10). These data suggest that statins do not have the ability to induce PPARγ-mediated

transcription when PPARγ protein levels are low, thus suggesting that PPARγ’s

transcriptional activity is not a key “player” in PTEN induction after statin stimulation.

Based on these results, Model C might be the most accurate representation of statin and

SREBP upregulation of PTEN expression in both CS and breast cancer patients (Fig.

3.3C).

Rosiglitazone does not inhibit SREBP induction of PPARγ-mediated transcription

It has been previously suggested that SREBP has the ability to induce the

production of natural PPARγ ligands(55). This indicates that SREBP’s ability to induce

both PPARγ protein and its ligand may be a mechanism for the increase in PPARγ’s transcriptional activity. Our above data demonstrate that statins can antagonize this pathway, potentially by interfering with PPARγ’s natural ligands. To test this hypothesis, we utilized Rosiglitazone, a synthetic but documented PPARγ ligand. We would expect

Rosiglitazone to signal down the same pathway as the natural PPARγ ligands and to induce PPARγ-mediated transcription, while not hindering ALLN’s induction. Therefore, we stimulated MCF-7 cells with Rosiglitazone, ALLN or the combination of

Rosiglitazone and ALLN, and analyzed PPARγ’s transcriptional activity. Rosiglitazone induced ~120-fold increase in luciferase activity, while ALLN induced ~160-fold increase in luciferase activity over EtOH-treated cells (Fig. 3.6). Interestingly, the combination of Rosiglitazone and ALLN (~150-fold) did not inhibit ALLN signaling as previously observed with the four statins (Fig. 3.5A). This suggests that statins hinder natural PPARγ ligand signaling. Additionally, these data confirm that statins do not induce PPARγ-mediated transcription, but rather, regulate a separate PPARγ protein activity to induce PTEN expression as illustrated in Model C (Fig. 3.3C).

Figure 3.6 SREBP signaling not inhibited by Rosiglitazone.

MCF-7 cells were transfected with PPRE-Luc and either empty vector (-; gray bars), wild-type PPARg (WT; black bars) or dominant-negative PPARg (DN; white bars). 12 hrs later cells were stimulated with 30 µM Rosiglitazone (Rosi), 15 µM ALLN alone or in Rosi/ALLN combination for 48 hrs and analyzed for luciferase activity as described.

Each bar represents a mean ± SEM of three individual experiments.

Statins upregulate PTEN transcription through an unknown transcription factor located between -854 and -791

Our results, consistent with our proposed Model C (Fig. 3.3C), show that statin stimulation increases PTEN transcription in a PPARγ transcriptional activation- independent mechanism, suggesting the involvement of another transcription factor(s).

To examine this in greater detail, we performed transcriptional reporter assays using serial deletions of the PTEN promoter. Seven PTEN promoter constructs were cloned into pGL3-Basic vectors: -1334 to 0, -1158 to 0, -1026 to 0, -893 to 0, -601 to 0, -453 to 0 and

-203 to 0; and subsequently transfected into MCF-7 cells. Twelve hrs after transfection,

MCF-7 cells were treated with either EtOH or one of the statins. Luciferase activity was measured and the results indicated that only one construct, that containing -893 to 0, could significantly induce PTEN transcription (~3.5-fold) over EtOH-treated cells (data not shown). Using this as a starting point, we constructed six more serial deletion vectors

(Fig. 3.7A). These vectors were transfected into MCF-7 cells followed by statin treatment. Only the construct containing the PTEN promoter region between -893 and -

601 could be significantly induced after Lovastatin (~2.5-fold), Simvastatin (~6.5-fold),

Pravastatin (~6.5-fold), and Fluvastatin (~4.0-fold) stimulation (Fig. 3.7B).

Figure 3.7 Statins induce PTEN transcription within the minimal promoter.

A) Illustration of serial deletion constructs of the PTEN promoter. B) MCF-7 cells were transfected with serial dilution constructs of PTEN promoter. Cells were then stimulated with vehicle, 3 µM Lovastatin, 3 µM Simvastatin, 30 µM Pravastatin, or 18 µM

Fluvastatin for 48 hrs and analyzed for luciferase activity as described. Each bar represents a mean ± SEM of three individual experiments. C) WT PTEN promoter -893 to -601 (lanes 1-4), -893 to -755 (lanes 5-8), and -769 to -608 (lanes 9-12) was radiolabeled and used as probes for EMSA analysis. Each promoter probe was either incubated in the presence (Bd) or absence (Ng) of nuclear protein and subsequently

competed with a nonspecific (Nn) or specific (Sp) competitor. A representative blot from three individual experiments is displayed. D) WT PTEN promoter -893 to -791 (lanes 1-

4), -893 to -834 (lanes 5-8), -854 to -791 (lanes 9-12), -854 to -755 (lanes 13-16), and -

811 to -755 (lanes 17-20) was radiolabeled and used as probes for EMSA analysis.

Nuclear protein was either not incubated with the probe (Ng), or incubated with the probe

(Bd) to test binding. Competition assays were performed with a nonspecific competitor

(Nn), and unlabeled WT promoter probe (Sp). A representative blot from three individual experiments is displayed.

Our PTEN promoter reporter assays demonstrated that statin stimulation induced transcription from a site between -893 and -601, which is interesting since there are no known transcription factors that bind to this region. To more accurately define the nucleotides of interest within this region, we utilized three different PTEN promoter probes. As expected, no retarded bands were observed during EMSA when nuclear protein was not added to the system (Ng). Protein binding (Bd) was observed utilizing the

-893 to -601 probe (Fig. 3.7C, 893-601), while the addition of a nonspecific competitor

(Nn) did not alter the binding and a specific competitor (Sp) did. This probe was subsequently divided into two additional probes: -893 to -755 and -769 to -608. The -893 to -755 probe demonstrated specific protein binding (Fig. 3.7C, 893-755), while the more

3’ probe did not reveal any protein binding (Fig. 3.7C, 769-609). Interestingly, our laboratory, while studying patient promoter variants that affect protein translation, identified this region as a target for the binding of a yet unidentified transcription factor(s)(9). In order to further dissect out this region of binding, we constructed five more serial deletion probes. Fig. 3.7D shows specific protein binding for three of the five probes with the smallest region of interest on the PTEN promoter that the statins regulate being between -854 and -791. Overall, therefore, our data suggest that statins upregulate

PTEN transcription by regulating PPARγ protein activity, which subsequently induces an as yet unknown transcription factor that binds to the PTEN promoter between -854 and -

791. Additionally, we were also able to demonstrate, for the first time, that SREBP can also induce PTEN transcription through upregulation of PPARγ-mediated transcription.

3.4 Discussion

PTEN is a constitutively active dual specificity phosphatase tumor suppressor. As

such, regulation of activity is determined by protein level. We have demonstrated that

statins, more specifically Lovastatin, Simvastatin, Pravastatin, and Fluvastatin, induce

PTEN transcription through regulating PPARγ protein activity, rather then its

transcriptional activity. Additionally, we subsequently isolated a small region of the

PTEN promoter, -854 to -791 that the statins regulate to induce PTEN expression.

Other groups have shown that statins can increase PPARγ mRNA and

subsequently its protein expression(56, 57). Our current study indicates that statins may

secondarily induce PPARγ transcriptional activity independent of PTEN regulation. We

demonstrated that the statins induced PTEN expression in the presence of endogenous

PPARγ protein levels (Fig. 3.1 and 3.2A), however, additional exogenous PPARγ was required for the statins to induce PPARγ’s transcriptional activity (Fig. 3.2B). The addition of exogenous PPARγ was essential because MCF-7 cells express low levels of

PPARγ (Fig. 3.4A). This indicated that PPARγ protein is the limiting factor in statin regulation of its transcriptional activity, but not for upregulation of PTEN expression.

Moreover, we established that in the presence of WT-PPARγ, but not DN-PPARγ, an

induction of PPARγ-mediated transcription could be observed after statin stimulation

(Fig. 3.2B). This further indicates that the statins’ ability to induce PPARγ transcriptional

activity is a specific effect; however, it is also clear from our work that it is not the

primary pathway for PTEN induction.

We initially proposed that statins induced PPARγ-mediated transcription to increase PTEN levels, however, this hypothesis was incorrect. These results were unexpected because we previously demonstrated that PPARγ protein expression is necessary to observe the induction of PTEN expression(51). Another protein that the statins have been connected with and is involved in regulating lipid metabolism, is

SREBP(54) which has been shown to be involved in PPARγ’s protein production and transcriptional activity(55). This raises the possibility that statins, PPARγ, and SREBP may all play a role in PTEN expression. Our results indicate that both the statins and

SREBP can signal through PPARγ, however, they do so differently and independent of each other. SREBP principally utilized PPARγ’s transcriptional activity, while statins primarily signal through a pathway independent of PPARγ’s transcriptional activity.

These results are similar to Ravid and colleagues’ study that showed ALLN inhibited

HMG-CoA reductase degradation(58). The combination of statin treatment with increased SREBP actually antagonized the two pathways (Fig. 3.5B). In contrast,

Mascaro and colleagues observed a Fluvastatin/ALLN synergistic effect on transcriptional activity in CHO cells(59). Our data indicate that SREBP can induce PTEN expression (Fig. 3.4A), suggesting that its upregulation of PPARγ-mediated transcription induces PTEN transcription. Based on these results, SREBP agonists may aid patients who demonstrate a decrease in PTEN but do not have an isolated PTEN mutation in the open reading frame; unfortunately, a clinically relevant SREBP agonist has yet to be established. However, one needs to be conscious of the idea that the combination of statins and a SREBP agonist may actually hinder the effect of each individual treatment.

Overall, our data strongly suggest that statins regulate a transcription factor(s), besides PPARγ, to induce PTEN transcription. We have demonstrated that Lovastatin,

Simvastatin, Pravastatin, and Fluvastatin upregulate PTEN transcription in the context of the PTEN promoter defined by a region flanking nucleotides -893 and -601 (Fig. 3.7B) and that an unknown transcription factor(s) bind(s) between -854 and -791 (Fig. 3.7D).

Only two transcription factors are predicted by multiple prediction software programs to bind at this region: Sp1 (TESS(60) and Alibaba(61)) and c-Myb (TESS and

TFSEARCH(62)). Sp1 is currently thought to be a putative PTEN transcription factor, due to the full-length PTEN promoter being very GC rich, however, empiric research has yet to concretely show that it binds to any particular region of the promoter or has the ability to regulate its transcription(6, 17). c-Myb has been shown to be upregulated within tumors when PTEN expression was decreased(63), indicating that it may be acting as a

PTEN transcriptional repressor, however, the pathway connecting the two has yet to be determined. Thus, both Sp1 and c-Myb may be postulated to be regulators of PTEN expression; however, more in depth studies are necessary to determine the identity of this novel PTEN transcription factor and are beyond the scope of this paper.

Our data reinforce the importance and benefit that agonists which upregulate

PTEN transcription may be useful to a subset of breast cancer patients. This suggests that a therapeutic tool that can regulate PTEN’s transcription would be highly effective within the subset of patients that have an identified nucleotide alteration within the PTEN promoter or in patients in which a PTEN mutation has yet to be identified. This approach would also be useful where hemizygous deletions occur, as in sporadic neoplasias, in increasing PTEN protein levels. This study indicates that Lovastatin, Simvastatin,

Pravastatin and Fluvastatin as well as a SREBP agonist would be germane to this cohort of patients. However, despite these encouraging results, we must be aware that these potential therapies may theoretically harm patients with germline intragenic PTEN mutations or those with neoplasias with somatic intragenic mutations by raising levels of mutant, as well as wildtype, protein.

CHAPTER 4

COWDEN SYNDROME PATIENTS WITH PTEN PROMOTER MUTATIONS

DEMONSTRATE ABNORMAL PROTEIN

4.1 Introduction

Germline mutations in PTEN (phosphatase and tensin homolog deleted on chromosome ten), a tumor suppressor gene on 10q23, occur in 85% of CS [MIM 158350] patients(4-6, 64, 65). This syndrome affects approximately 1 in 200,000 individuals, however, this is generally thought to be an underestimate(30). CS is characterized by hamartomas of multiple organs and increased risks of neoplasia. Patients diagnosed with

CS have a 25-50% lifetime risk of developing female breast cancer, while the risk for the general population is ~13%(29, 49). Thyroid cancer in CS patients, which tends to be follicular, has ~10% lifetime risk, compared to <1% in the general population.

Furthermore, there is a ~5-10% lifetime risk of endometrial cancer in CS patients compared to ~2-4% in the general population(6, 29). Proper recognition and diagnosis of

CS patients is crucial not only due to the increased risk of cancers observed in this syndrome, but also due to the morbidity and even mortality associated with its non- malignant features.

Germline PTEN mutations are also found in 65% of BRRS [MIM 153480] patients, characterized by macrocephaly, lipomatosis, hemangiomatosis and speckled penis(6, 8). In addition to BRRS, a number of other syndromes including PS [MIM

176920], PLS(30) and autism(66) share germline PTEN mutations as an etiology and have been classified as PHTS. It is recommended that they are all managed in a similar manner to CS if a pathogenic PTEN mutation is identified(5).

Prior to 2003, PTEN mutations had been identified in 80% of patients meeting strict diagnostic criteria for CS(2, 49). CS is believed to be mono-genic, therefore determining the cause of PTEN dysfunction in the remaining 20% of patients is vitally important to the practice of personalized genetic healthcare. Our laboratory began aggressively pursuing alternative mechanisms of PTEN inactivation including interrogating the mutation status of its own promoter. We have identified mutations in the

PTEN promoter in CS patients and have shown that ~10% of previously classified PTEN mutation negative patients have nucleotide variants within the full-length promoter(6).

Furthermore, these mutations resulted in both a decrease in PTEN protein expression and loss-of-function. In contrast to CS patients, no PTEN promoter mutations have been identified in BRRS patients to date, although large deletions involving PTEN were identified in approximately 10% of mutation negative BRRS patients(6). Interestingly, in the initial study, eight of nine (89%) patients with PTEN promoter mutations had breast cancer, although the overall number of organs involved was generally low in these patients suggesting these mutations may preferentially confer very high penetrance for breast cancer(6).

Previous PTEN promoter investigations, including our own, have focused on areas within known transcription factor binding sites. However, we have identified variants of unknown significance (VUS) 3` of any known cis-acting elements. Based upon our previous work(17, 23, 67), we would predict that these VUSs might be pathogenic. Given the dilemma faced by the clinical cancer geneticist and genetic counselor when counseling patients with a VUS, a deeper understanding of the molecular consequences of these promoter VUSs would have important implications for patient care. In this study, we investigated both transcriptional and translational downstream effects of PTEN promoter VUSs in patients with CS. Our data demonstrate that certain

PTEN promoter VUSs in patients with CS result in decreased PTEN expression through dysfunctional translation, rather than through altered transcription.

4.2 Materials and Methods

Patient recruitment

Human subjects were recruited from multiple institutions throughout the United

States. All samples were acquired with informed consent in accordance with approved protocol by the human subjects protection committees of their respective institutions.

Cowden syndrome patients used in this study were classified in accordance with both the

National Comprehensive Cancer Network and the International Cowden Consortium operational diagnostic criteria. 186 normal healthy individuals were analyzed for controlled comparison studies. Patients are approximately 85% white, ~5% black and

10% other (including Hispanic, Orientals, etc).

DNA Isolation and Promoter Mutation Analysis

Normal control and patient genomic DNA was isolated by the Genomic Medicine

Biorepository of the Cleveland Clinic Genomic Medicine Institute (see URL on Web

Resources for more information). Primers were designed to amplify the full PTEN

promoter between -1389 and -715 (F: 5`-GCGTGGTCACCTGGTCCTTT-3` R: 5`-

GCTGCTCACAGGCGCTGA-3`) and DNA was PCR amplified in 20 ul reactions using

HotStar and Q Solution (Qiagen). The PCR conditions consisted of 30 cycles at the

annealing temperature of 55°C. PCR products were treated with exonuclease I (New

England Biolabs, Ipswich, MA) and shrimp alkaline phosphatase (USB, Cleveland, OH)

and were analyzed on both strands by direct sequencing (ABI 3730xl DNA Analyzer,

Genomics Core Facility, Lerner Research Institute, Cleveland Clinic). Variants were

detected by direct analysis compared with normal control sequence through Lasergene

software (DNASTAR, Madison, WI).

Cell culture

The MCF-7 breast cancer and HeLa cell lines were maintained at 37°C with 5%

CO2 in DMEM containing 10% FBS and 100 units/ml each penicillin and streptomycin.

Clonal lymphoblastoid cell lines (LBCL) were generated by the Genomic Medicine

Biorepository from CS patients and normal healthy controls (see URL on Web Resources for more information). LBCLs were cultured in RPMI medium supplemented with 20%

FBS and 100 units/mL penicillin and streptomycin.

Protein Isolation

Total protein from LBCLs was harvested using M-PER (Pierce; Rockford, IL)

lysis buffer containing PMSF (0.75mg/ml), benzamidine hydrochloride (0.5 mg/ml),

leupeptin (2µg/ml), aprotinin (2 µg/ml), pepstatin (2 µg/ml), β-glycerophosphate (10

mM), NaOV (0.2 mM), and NaF (25mM). Cells were incubated at room temperature with

lysis buffer for 1 min before harvesting. Samples were centrifuged at 16,000 X g for 10

min at 4ºC to remove cellular debris. The resulting supernatant was stored at -80°C.

Nuclear protein was extracted according to Pierce’s isolation protocol (Madison, WI). In

brief, cells were collected by scraping with PBS and washed several times. Cell pellets

were incubated with the appropriate amount of Cer I buffer on ice to isolate the

cytoplasmic fraction. The remaining extract was incubated with the appropriate amount

of Ner I buffer on ice with several agitation steps to isolate the nuclear fraction. The

resultant cytoplasmic and nuclear extracts were stored at -80°C. Protein concentration

was determined using the bicotinic method using BSA as a standard (56).

Electromobility Shift Assay (EMSA)

To study the VUSs in a heterozygous state, the PTEN promoter sequence (-893 to

-755) was isolated through PCR amplification from either normal genomic DNA or from genomic DNA obtained from a CS patient that has an identified variant. The DNA was

PCR amplified with 30 cycles at the annealing temperature of 55°C in 20 ul reactions using HotStar and Q Solution (Qiagen). (F: 5`-ATGCGCTGCGGCAGGATAC-3`; R: 5`-

CTCATCTCCCTCGCCTGA-3`). To study the VUSs in a homozygous state, the above

PCR products were cloned into TOPO-TA vectors and product sequences were verified 83

by direct DNA sequencing (ABI 3730xl DNA Analyzer). Nested PCR was subsequently

performed with the above primers to isolate only the PTEN VUSs. Each of the above products was radiolabeled with 32P-γATP via T4 kinase. To examine DNA-protein

interaction 1 ng of radiolabeled probe was incubated with 2 µg of nuclear protein extract

for 20 min at room temperature with binding buffer containing 10 mM HEPES (pH 7.5),

2.5 mM MgCl2, 50 mM NaCl, 0.5 mM DTT, 4% glycerol, 1 µg/mL BSA, and 2 µg poly

dI/dC. Unlabeled probe in 5X molar excess was used as the specific competitor, while a

random oligonucleotide sequence was used as the nonspecific competitor. DNA:protein

complexes were resolved on a 4% nondenaturing PAGE gel at 150 V for 3.5 hrs at 4°C

and visualized using a Phospho-Imager (Amersham Biosciences, Piscataway, NJ).

Reporter Assay

PTEN (–893 to –1) was PCR amplified from normal genomic DNA or CS patient

genomic DNA (F: 5` GCGTGGTCACCTGGTCCTTT 3`; R: 5`

GCTGCTCACAGGCGCTGA 3`) and subsequently cloned into a TOPO-TA vector.

DNA was PCR amplified using 55°C as the annealing temperature for 30 cycles. All PCR

amplification products were verified by direct DNA sequencing (ABI 3730xl DNA

Analyzing), and positive clones were subcloned into a pGL3.1-Basic vector (Promega,

Madison, WI). In the event that positive variant clones were not obtained, site-directed

mutagenesis was performed, as described in manufacture’s protocol (GeneTailor Site-

Directed Mutagenesis System; Invitrogen). Site-directed mutagenesis was performed on

the WT PTEN promoter in a TOPO-TA vector.

To determine promoter activity, 6-well plates of MCF-7 or HeLa cells were co-

transfected with 1 µg/well of a PGL3-PTEN construct, and 50 ng/well Renilla luciferase

control plasmid with 3µl/well of FuGene (Roche,) as described by the manufacture. After

48 hrs cells were harvested with 1X passive luciferase lysis buffer (Dual-Luciferase

Reporter Assay System; Promega) and analyzed on a luminometer (LMax 11384;

Molecular Devices) using Renilla luciferase as an internal transfection control.

Western Blot

15 µg of protein was prepared by the Laemelli method(36), separated on a 10%

SDS-PAGE gel, and electrophoretically transferred onto nitrocellulose. Equal protein

loading between conditions was confirmed by staining with Ponceau S solution. Non-

specific binding was blocked by incubating the nitrocellulose blots with 5% milk in TBS-

T (100mM Tris, pH 7.5, 1M NaCL, 1% Tween-20) for 1 hr at room temperature. Blots

were then incubated with the primary antibody, either anti-PTEN (Cascade Bioscience,

Waltham, MA) or anti-actin (Sigma, St Louis, MO) at a dilution of 1:1000 in 3% BSA

for 2 hrs at room temperature. Following the primary incubation the blots were washed

with TBS-T for 1 hr with frequent changes of buffer. Blots were then incubated with the

appropriate secondary antibody conjugated to horseradish peroxidase (Promega;

Madison, WI) at a dilution of 1:2500 in 5% milk overnight at 4°C, and washed with TBS-

T for 1 hr. Protein bands were visualized using enhanced chemiluminescence as

described by the manufacturer (Amersham Pharmacia Corp; Piscataway, NJ).

RT-PCR and Real-Time PCR

HeLa and MCF-7 cells transfected with the above PGL3_PTEN -893 to -1 construct and LBCLs were collected and subsequently washed three times with PBS through centrifugation. Total RNA was extracted from cells following the Gentra

Versagene RNA Purification System Protocol (Minneapolis, MN) and then converted to cDNA by SSIIRT after DNase treatment. The resultant cDNA was subjected to multiplex

PCR amplification using primers specific to luciferase (F: 5’

TCAAAGAGGCGAACTGTGTG 3’ and R: 5’ GGTGTTGGAGCAAGATGGAT 3’),

PTEN exon 3 and exon 5 (F: 5’ TGGATTCAAAGCATAAAAACCA 3’; R: 5’

AAAAGGATATTGTGCAACTCTGC 3’) and β-actin (Quantum RNA β-actin; Ambion

INC; Austin, TX). Primers were allowed to anneal at 55°C for 28 cycles. The products from the PCR reactions were run on a 1% agarose gel containing ethidum bromide and visualized under a UV light.

Real time PCR was carried out using the ABI 7500 real time PCR system

(ABI/Perkin Elmer, Foster City, CA) using a SYBR green-based assay as previously described(68). Primers were designed to amplify cDNA incorporating a portion of the

PTEN transcript encoded by exons 7 and 8 (7F: 5’ CCACAAACAGAACAAGATG 3’ and 8R: 5’ CTGGTCCTGGTATGAAGAAT 3’). Primers amplifying a portion of

GAPDH were used as the control (F: 5’ CCATCTTCCAGGAGCGAGA 3’ and R: 5’

AAATGAGCCCCAGCCTTCT 3’). The thermal cycling conditions were 50oC for 2 min and 95oC for 10 min, followed by 40 cycles of denaturing at 95oC for 15 sec and annealing and extension at 60oC for 1 min. All the reactions were performed in triplicate, and the comparative CT method was used for the quantification of the expression for each 86

segment, by use of GAPDH as a normalization control. Each PCR reaction generated only the expected amplicon as shown by the melting temperature profiles of the final products and by gel electrophoresis.

Statistical Analysis

Statistical analysis was done using Student’s t test. Data are mean ± SD of three independent experiments and normalized to a control. P<0.05 is considered significant.

4.3 Results

PTEN promoter VUSs demonstrate similar transcription factor binding

Despite the clear significance of PTEN dysfunction in the development of many types of cancers, the mechanisms that regulate PTEN’s promoter and 5’-UTR largely remain unknown. Figure 4.1 illustrates that the generally recognized full-length PTEN promoter is localized to position -1344 to -745, and the minimal promoter as -958 to -

821, with +1 representing the ATG site(19). While scanning the full-length PTEN promoter for nucleotide variants, we identified five VUSs (-861G/T; -853C/G; -834C/T; -

798G/C; -764G/A) in five unrelated patients with a clinical diagnosis of CS, when compared to 186 normal healthy controls. All five CS patients had benign and/or malignant breast disease and importantly 3/5 (60%) had two component malignancies associated with CS (Table 4.1). Additionally, all five of the VUSs used in this study are located 3` of any known PTEN transcription factor-binding site (Fig 4.1). Previous data from our laboratory indicate that mutations within regulatory elements or transcription

factor binding sites interfere with normal transcriptional activity(17, 23, 67). However, these five VUSs are not predicted to alter any known cis-acting regulatory elements therefore determining the functional significance of these variants is even more difficult to ascertain.

Malignant Benign Breast Follicular Endometrial Mutation Breast Cancer Neoplasm Thyroid Cancer Cancer

-861G/T No Yes Yes Yes -853C/G Yes No Yes No -834C/T Yes No No No -798G/C No Yes Yes Yes -764G/A No Yes No Yes

Table 4.1 Increase frequency in Cowden syndrome patients with PTEN promoter variants.

Figure 4.1 Schematic diagram of the PTEN promoter.

PTEN’s full-length promoter lies between -1344 and -745 (white bar), while the minimal promoter lies between -958 and -821 (gray bar). Within this region, five nucleotide variants (black triangles), -861G/T, -853C/G, -834C/T, -798G/C and -764G/A, have been identified in a subset of classic CS patients. The nearest 5’ transcription factors are EGR1 (dotted bar) and Sp1 (black bar).

To determine what role these five VUSs may play in PTEN regulation, we

performed electromobility shift assays (EMSAs) to find out if nuclear protein has the

ability to associate with the promoter at this region. Interestingly, we found that nuclear

protein does bind to the WT PTEN promoter from -893 to -755, suggesting there is a yet to be identified transcription factor binding site in this region. We continued further to determine if the nucleotide variants resulted in altered DNA:protein complex formation when compared to WT PTEN promoter. EMSAs for each VUS were performed in either a homozygous state, only the variant allele present, or in a heterozygous state, where both the WT and variant allele were present. This allowed us to determine if the variant allele has the ability to inhibit the WT allele’s function. As expected, no DNA:protein complex was observed when nuclear protein was not added to the reaction mixture (Fig 4.2A, Ng, lane 1) and nuclear protein does bind to WT PTEN promoter from -893 to -755 (Fig

4.2A, Bd, lane 2). Unexpectedly, we observed that nuclear protein was also able to bind the PTEN promoter VUSs in both a homozygous and heterozygous state (Fig. 4.1A, Bd, lanes 4, 6, 8, 10 and 12). Furthermore, these interactions did not differ significantly between the WT and VUS PTEN promoter probes (Fig 4.2B). In addition, the WT PTEN

promoter DNA:protein complex could be competed with a cold unlabeled VUS probe

(Fig. 4.2C). Taken together, these data suggest that the DNA:protein interaction at this

site is not affected by these specific promoter VUSs.

Figure 4.2 EMSA analysis probed with radiolabelled PTEN promoter from -893 to -755, which is either WT or variant.

A) Each promoter probe was either incubated in the presence (Bd) or absence (Ng) of nuclear protein. The WT PTEN promoter (lanes 1,2), -861G/T (lanes 3,4), -853C/G

(lanes 5,6), -834C/T (lanes 7,8), -798G/C (lanes 9,10) and -764G/A (lanes 11,12)

demonstrate nuclear protein binding. A representative blot from three individual

experiments is displayed. B) Quantification of nuclear protein binding to the PTEN

promoter confirms insignificant differences between WT (lane 1) and the five variants

(lanes 2-6, p>0.050). EMSA results are depicted as fold-change compared to WT PTEN

promoter and are shown in graphical format. (Student’s t-test) C) WT PTEN promoter from -893 to -755 was radiolabeled. Nuclear protein was either not incubated with the probe (Ng), or incubated with the probe (Bd) to test binding. Competition assays were performed with a nonspecific competitor (Nn), unlabeled WT promoter probe (Sp) and cold PTEN promoter variant probes (lane 5-9). A representative blot from three individual experiments is displayed.

PTEN promoter reporter assays demonstrate VUSs significantly inhibit luciferase activity

The above data suggest that transcription factor binding at position -893 to -755 is not altered in patients who harbor VUSs within this region, and that formation of this complex may not directly contribute to disease pathogenesis. In order to gain more insight regarding the potential mechanism(s) of PTEN dysfunction manifested by these variants, we examined the ability of each VUS to promote PTEN transcription via reporter assays. We observed that the WT PTEN (position -893 to -1) had a basal level of luciferase activity (Fig 4.3, WT). Two of the patient promoter VUSs, -834C/T and -

798G/C, had similar luciferase read-outs (Fig. 4.3). In contrast, three of the VUSs showed an inhibition in luciferase activity when compared to the WT PTEN construct. We found that the -861G/T and -764G/A VUSs had the greatest inhibition of luciferase activity, approximately 50% less then that of the WT construct (Fig. 4.3; p<0.001). Additionally, the -853C/G VUS resulted in a 40% decrease in luciferase activity, compared to WT

PTEN (Fig. 4.3; p<0.001).

Figure 4.3 Luciferase activity of PTEN 5’UTR altered in three VUSs

MCF-7 cells were transfected with PGL3_PTEN 5’UTR from -893 to -1 that is either WT or variant, as described in the Methods section. After 48 hrs of treatment, cells were harvested and luciferase activity was measured. Each bar represents a mean ± SEM of three individual experiments. *p<0.001 (Student’s t-test)

Promoter VUSs do not inhibit PTEN transcription

The above experiment indicates that WT PTEN induces more luciferase activity

compared to three of the patient-derived promoter VUSs, specifically the -861G/T, -764

G/A and -853C/G variants. However, this experiment does not indicate if this effect is

due to a decrease in transcription or an inhibition of protein translation. In order to

differentiate the two scenarios, we analyzed luciferase mRNA levels in the reporter assay

samples described above. If the promoter VUSs are affecting transcription, we would

expect to see changes in luciferase mRNA levels in those VUSs that resulted in significant decreases in luciferase activity. Unexpectedly, we found that luciferase mRNA levels were equally expressed in the cells transfected with the WT PTEN

construct compared to cells transfected with the patient-derived VUS promoters (Fig

4.4A), suggesting that transcription efficiency and mRNA stability are not compromised

in these five VUSs.

We next examined the VUSs ex-vivo by assessing PTEN mRNA levels using total

RNA isolated from LBCLs from patients with promoter VUSs. We were able to assess

this in three cell lines derived from the patients with the following VUSs: -861G/T, -

853C/G, and -798G/C. We were unable to obtain LBCLs from the patients with the

remaining two VUSs (-834C/T and -764G/A). By semi-quantitative RT-PCR, we

demonstrated that there was equivalent PTEN mRNA expression in LBCLs harboring promoter VUSs compared to those obtained from normal, healthy controls (Fig. 4.4B, p>0.05). To confirm these results we performed a quantitative real time PCR assay.

Similarly, no differences were observed between PTEN mRNA levels in normal, healthy controls compared to patient-derived promoter VUSs (Fig 4.4C, p>0.05).

Figure 4.4 WT and variant PTEN promoters demonstrate equal mRNA expression

A) HeLa or MCF-7 cells were transfected with a PGL3_PTEN -893 to -1 construct that was either WT or contained a VUS. 48 hrs after transfection cells were harvested and total RNA was extracted. Luciferase (top panel) and actin (bottom panel) mRNA levels were measured by RT-PCR. p>0.050 Represents a mean ± SEM of three individual experiments. B) LBCLs from either normal healthy controls that are PTEN WT (lane 1-3) or patient-derived with a promoter VUS (-861G/T lane 4, -853C/G lane 5, and -798G/C lane 6) were harvested and total RNA was extracted. PTEN (bottom band) and actin (top band) mRNA levels were measured by RT-PCR. p>0.050 Represents a mean ± SEM of three individual experiments. C) LBCLs from either normal healthy controls that are

PTEN WT (bars 1-3) or patient-derived with a promoter VUS (-861G/T bar 4, -853C/G bar 5, and -798G/C bar 6) were harvested and total RNA was extracted. Real time measured PTEN and GAPDH mRNA levels. Quantification of PTEN mRNA normalized to GAPDH levels. Real time RT-PCR results are depicted as ∆Ct changes and are shown in graphical format. Each bar represents a mean ± SEM of three individual experiments. p>0.050 (Student’s t-test)

PTEN mRNA secondary structure is altered in VUSs versus PTEN WT promoter

As the above data suggest, these PTEN VUSs do not result in altered transcription. Therefore, we next focused on whether they result in abnormal translation by studying the PTEN mRNA transcript in more detail. Several laboratories have previously suggested a number of putative PTEN transcriptional start sites between -1031 and -93(19, 20, 26, 33), however, comparison of the human and mouse PTEN cDNA sequences suggest that transcript begins around -925(20). In agreement with these results, we performed PTEN RT-PCR from -869 to exon 9 and verified that all of the five VUSs are included in the resulting transcript (data not shown). Therefore, we hypothesized that the inclusion of these nucleotide variants within the transcript causes an alteration of the normal mRNA secondary structure, and consequently an inhibiting protein translation. To determine if the mRNA secondary structure is different in the VUSs compared to WT

PTEN, we utilized the MFOLD software program(69). We analyzed the PTEN 5’UTR from position -893 to -1, using both the WT sequence and sequences containing each of the VUSs. Several potential secondary structures for each sequence were predicted, with the most stable structures illustrated in Figure 4.5.

Figure 4.5 MFOLD-predicted secondary structures resulting from the five VUS in

Cowden syndrome patients

The most stable mRNA secondary structures predicted by MFOLD are illustrated here:

WT (top left panel), -861G/T (top middle panel), -853C/G (top right panel), -843C/T

(bottom left panel), -798G/C (bottom middle panel) and -764G/A (bottom right panel).

The major secondary structure predicted for the WT PTEN promoter is Y shaped with multi-loops (Fig 4.5, WT). This shape is very consistent with that predicted for the -

834C/T and -798G/C (Fig 4.5) VUSs. In contrast, the -861G/T, -853C/G and -764G/A

VUSs have different predicted secondary structures compared to the WT PTEN 5’UTR.

While the -853C/G VUS maintains the general Y shape, the loop structures are altered to

create a new branching arrangement (Fig 4.5, -853C/G). Similarly, the -764G/A VUS

maintains the Y shape that is similar to the WT 5’UTR; however, it has lost almost all of

its loops creating more of a stick-like structure (Fig 4.5, -764G/A). Finally, the -861G/T

VUS is structurally the most different from WT PTEN (Fig 4.5, -861G/T). The general Y

shape appearance has been lost and replaced with a detailed, intricate looping and

branching configuration. These structures suggest that PTEN translation may be altered

or inhibited in patients with these promoter VUSs, particularly those that have the

greatest deviation from the WT structure, such as -861G/T.

Altered PTEN protein expression in select VUSs

In order to determine if these patients do demonstrate altered protein translation

due to nucleotide variants within the promoter region, we examined PTEN protein

expression from the previously mentioned patient-derived LBCLs. We found that protein

isolated from normal healthy controls had similar PTEN protein expression (Fig 4.6A/B,

lanes 1-3). Comparable to our control samples, normal PTEN protein expression was

observed from patient-derived LBCLs with VUS -798G/C (Fig 4.6A/B, lane 6). Protein

lysate derived from patient LBCLs with the -853C/G VUS had a ~15% decrease in PTEN

expression (p=0.046; Fig 4.6A/B, lane 5), while cells derived from VUS -861G/T had the

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largest decrease in PTEN levels at ~40% of control protein (p<0.001; Fig 4.6A/B, lane 4).

Interestingly, these protein results can be seen as concurrent with the structural alterations

observed with the MFOLD software. The secondary structure of VUS -798G/C 5’UTR

does not predict a large change compared to WT PTEN therefore one would predict that

PTEN protein levels would be comparable to WT controls as we observed here. In contrast, the secondary structure predicted with the -853C/G 5’UTR VUS predicted some alterations, which correlate with the slight decrease in PTEN protein levels observed in this patient. Finally, the VUS promoter with the greatest structural change, -861G/T, demonstrated the largest alteration in PTEN levels. Taken together, these data indicate that these predicted alterations to the secondary structures of the 5’UTR including the

PTEN promoter, consequent to the VUSs in our CS patients, inhibit normal translation of

PTEN.

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Figure 4.6 PTEN protein expression decreased in promoter variants with the greatest mRNA secondary structure alterations

A) LBCLs from either normal healthy controls that are PTEN WT (lane 1-3) or patient- derived with a promoter VUS (-861G/T lane 4, -853C/G lane 5, and -798G/C lane 6) were harvested and total protein was extracted. PTEN (top panel) and actin (bottom panel) protein levels were measured by western blot analysis. B) Quantification of PTEN protein normalized to actin levels. Western blot results are depicted as fold-change and are shown in graphical format: -861G/T (p<0.001), -853C/G (p=0.046) and -798G/C

(p>0.050). Each bar represents a mean ± SEM of three individual experiments. *p<0.050

(Student’s t-test) 102

4.4 Discussion

Recently, the importance of gene regulation in the pathogenesis of hereditary

cancer predisposition syndromes has been advanced though the interrogation of promoter

variation as a mechanism of disease development. PTEN promoter variants and their

consequences have only been minimally studied, however promoters within a few select

genes, such as baculoviral IAP repeat-containing 5 (BIRC5 also known as survivin), have

been examined more extensively(70). Yet the majority of these studies have looked

specifically at known regulatory regions and/or consensus sites within the promoter of

interest. We hypothesized that novel nontraditional regulatory mechanisms within the

PTEN promoter play an important role in gene regulation. This indicates that variants

within unknown regulatory elements may be as significant as those in known cis-acting

regions. To test this hypothesis, we studied five VUSs within the PTEN promoter that are

not within a known cis-acting element. The culmination of our data reveal that protein

translation is altered within a subset of patients with CS who lack traditional exonic or

splice-site PTEN mutations but harbor PTEN promoter variants, particularly those

resulting in large mRNA structural changes compared to WT PTEN mRNA. These data

also demonstrate abnormal protein translation as a novel mechanism of CS pathogenesis.

To date, analysis of PTEN’s promoter has identified eight regulatory factors that

have been implicated in modulating PTEN’s transcription: Egr-1(71), NF-κB(26), Sp1(6,

33), CBF-1(24), and p53(20), USF1(23), PPARγ(21, 50, 51) and c-Jun(56) (Fig 4.1). The five VUSs (-861G/T; -853C/G; -834C/T; -798G/C; -764G/A) used in our studies reside in the full-length PTEN promoter region between -893 and -755, but lie more 3’ of any of 103

these known transcription factor-binding motifs. Our EMSA results, using PTEN -893 to

-755 as bait, demonstrated nuclear protein binding (Fig 4.2), thus suggesting that there

may be a novel transcription factor-binding site contained within this region. Several

potential transcription factors are anticipated to bind to this region, but only two were

predicted by multiple prediction software programs: Sp1 (TESS(60) and Alibaba(61))

and c-Myb (TESS and TFSEARCH (62)). Sp1 is currently thought to be a putative PTEN

transcription factor, due to the full-length PTEN promoter being very GC rich, however

research has yet to concretely show that it binds to any particular region of the promoter

or has the ability to regulate its transcription(6, 19). c-Myb has been shown to be

upregulated within tumors when PTEN expression was decreased(63), indicating that it

may be acting as a PTEN transcriptional repressor, however the pathway connecting the two has yet to be determined. Based on these prediction models, both Sp1 and c-Myb can be postulated as regulators of PTEN expression, however more in depth studies are necessary to determine the identity of this novel PTEN transcription factor.

In our initial EMSA results, we expected to observe a difference in this novel transcription factor’s ability to bind to the PTEN promoter, as previous data from our laboratory have shown that PTEN promoter alterations within the p53 (-1190 to -

1157)(17) and USF1 (-2237 and -2058)(23) binding sites inhibit both normal PTEN

mRNA expression and protein function. In contrast, data presented herein indicated that

protein-binding inhibition is not the primary mechanism of PTEN alterations (Fig 4.2).

Interestingly, our reporter assay results indicated that several of the VUSs had a decrease

in luciferase activity (Fig 4.3), while seemingly paradoxically, PTEN mRNA levels were

equally expressed relative to WT (Fig 4.4). This suggests that normal protein translation

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is disrupted by these alterations, while conventional mRNA transcription remains

unaffected. In 2001, Signori and colleagues observed a similar effect caused by a variant

located three nucleotides upstream of the ATG site, thus lying within the Kozak

consensus sequence of the 5’UTR within BRCA1(72). This nucleotide variant is thought

to have weakened the Kozak sequence enough to inhibit normal protein translation. In

contrast to Signori et al, the variants discussed in this publication did not immediately

indicate this mechanism and more intricate analyses were necessary.

PTEN has a number of putative transcription start sites, whose analysis reveals

five potential Kozak translational start sites. However none of these start sites perfectly

fit the mammalian Kozak consensus sequence, GCCRCCATGG, where the -3 and +4

positions are the most conserved(73). Analyses of these ATG sites, 5’ to 3’, indicate that

they would produce 28-amino acid (AA), 5AA, 4AA, 46AA and 403AA proteins,

respectively. PTEN is just one of many genes that has the potential to produce upstream

open reading frames (ORFs) and may not follow the “first ATG” rule. Moreover, this

tends to lead to leaky scanning by the 40S ribosome thus allowing for translation of one

or several of the 5’ ORFs. In this situation, the ribosomes do not fall off the transcript and

proceed to scan 3’ to the true ATG site, thus allowing for the production of the correct

mRNA. This suggests that PTEN’s long 5’UTR with potential ORFs and a weak Kozak

consensus sequence make it more prone to influences from nucleotide variants, which

can subsequently decrease its translation efficiency(74).

Another mechanism that can confound normal protein translation efficiency is

through aberrant mRNA secondary structures. To determine if modified PTEN mRNA structure was the cause of altered protein translation in our patients, we utilized the

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MFOLD software program to compare the WT PTEN promoter with the five VUSs. Our results demonstrate that some of these VUSs contribute to the mRNA structure, thus creating a significantly different confirmation compared to WT PTEN (Fig 4.5). This is important because normal mRNA primary and secondary structures are essential for accurate mRNA binding proteins ability to both bind and initiate, and subsequently affect protein translation(74). Recently, Saxena and colleagues identified an eleven base-pair deletion in MeCP2 103 nucleotides upstream of the ATG site(75). Through the use of mRNA structural prediction models, they concluded that this deletion disrupted normal protein translation without affecting MeCP2 transcription.

The nucleotide alterations isolated within both BRCA1 and MeCP2, which specifically inhibit normal translation and not gene transcription, are currently the only known reports to demonstrate that promoter VUSs can affect protein expression through such a mechanism. However, there have been a few recent examples of single nucleotide polymorphisms (SNPs) within protein coding regions that also alter the normal protein outcome through translation. In both of these examples, it is hypothesized that synonymous SNPs induce a structural changes in mRNA structure, which ultimately lead to the protein’s dysregulation. Kimchi-Sarfaty et al report specific MDR1 haplotypes, generated from silent SNPs, inhibit normal protein translation and subsequently its function. This is thought to occur through slowing down of ribosomal scanning at these specific codons(76, 77). Furthermore, Nackley and colleagues described three major haplotypes formed by four SNPs within the human catechol-O-methyltransferase

(COMT) gene who’s MFOLD predictions indicated that the haplotypes demonstrate altered mRNA secondary structures. More interestingly, a decrease in both protein

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expression and enzymatic activity was produced from a haplotype containing two

synonymous SNPs(78). These data suggest that nucleotide changes that may initially

seem insignificant can disrupt normal protein translation through alterations of mRNA

secondary structure.

As stated above, our reporter assay, MFOLD analysis, and western blot results are

all in agreement with regards to the -861C/G, -853C/G and -789G/C VUSs. Due to

unavailability, we were unable to directly study the -834C/T and -764G/A VUSs,

however, one can speculate on the likely outcome based on the above results. Our data

indicate that the -834C/T VUS, which displays no secondary structure differences

compared to WT PTEN mRNA, and had only a slight decrease in luciferase activity

would not have a decrease in PTEN protein expression. In contrast, one can hypothesize

that the -764G/A variant would have a significant decrease in PTEN protein expression.

Similar to the -861C/G VUS, the -764G/A VUS demonstrated a significant decrease in

luciferase activity and a large alteration in mRNA secondary structure, as predicted by

MFOLD, when compared to WT PTEN 5’UTR.

It is thought that the decrease in translation efficiency of these mRNA structures can be compensated for by more efficient translation through the regulation of the eukaryotic translation initiation factor (eIF4F) complex. This complex which is composed of eIF4A, eIF4B and eIF4H, is involved in unwinding mRNA secondary structures to induce protein translation(79). These data suggest one could potentially modulate one or several of these translation factors as a personal therapeutic target for patients with PTEN

promoter VUSs.

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In 2003, our laboratory was the first to demonstrate the pathogenicity of PTEN

promoter variants in patients with CS(6). These genetic alterations within PTEN’s

promoter appear to correlate with a high prevalence of breast cancer in this subset of

patients. In agreement with these previous data, the patients included in the current

analysis all harbor promoter VUSs and exhibit a high prevalence of breast neoplasia, but

in addition, follicular thyroid and endometrial cancer. All five patients with the PTEN

promoter VUSs interrogated in this study had developed breast tumors (Table 4.1). Two

of these five patients were diagnosed with breast cancer, while the remaining three

patients were diagnosed with benign breast neoplasms. In addition to breast cancer, 3/5

patients developed follicular thyroid cancer, and 3/5 patients had endometrial cancer

suggesting these VUSs are associated with neoplastic risk. All five patients ultimately

were diagnosed with at least one component malignancy, and 3/5 (60%) were diagnosed

with two component malignancies.

Despite their CS diagnosis, PTEN ORF and promoter mutation analysis, as well as deletion analysis, failed to definitively define the genetic etiology. Through more functional analysis of the VUSs located within the PTEN 5’UTR, we have now

elucidated this mechanism in 3/5 CS patients, each of whom harbors a previously

uncharacterized promoter VUS (-861G/T, -853C/G, and -764G/C). However, our data

indicate that aberrant protein translation is likely not the primary mechanism of CS in

patients with the -834C/T and -798G/C variants. One can speculate that within theses

patients, a specific haplotype may be playing a key role(67) or the PTEN protein function

is altered(80). This suggests that the region upstream of PTEN plays an important role, however the precise mechanism of these mutations remained to be elucidated.

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Through both this study and previous work from our laboratory, we have begun

paving the road to better understanding the function of the PTEN promoter and its role in

pathogenesis of CS. These data reinforce the importance of PTEN promoter nucleotide variations and their ability to lead to CS progression through protein translation inhibition. As discussed, CS patients with promoter mutations have a high prevalence of breast, thyroid and endometrial malignancies, and an understanding of the mechanism of

PTEN dysfunction in these patients, may lead to rational targeted therapies to treat or prevent malignancy. Our data suggests that a therapeutic tool that can regulate its transcription and/or translation, such as Lovastatin(51) or an eIF4F target, could be highly

effective within patients with germline nucleotide alterations within this region or in

sporadic tumors with somatic 5’-UTR VUS’s. Moreover, our data also reiterate the

importance of looking for variants within the PTEN promoter and even elsewhere in the

5’UTR in patients that have CS features yet do not have a detectable mutation within its

ORF. Furthermore, this knowledge should also be broadened to other diseases and their respective altered genes. Currently, promoters are rarely being studied in the clinical setting, therefore it is very likely that nucleotide changes will also be isolated within other genes and these patients may also benefit from personalized treatment given their promoter mutation status. Our observations should increase the frequency of germline

PTEN mutations in PHTS, thus increasing the sensitivity of molecular diagnosis, and hence, broadening those families amenable to predictive testing.

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CHAPTER 5

DISCUSSION

PTEN is a constitutively active, dual-specificity phosphatase, tumor suppressor.

As such, regulation of its activity is determined by its protein level. One mechanism of regulating PTEN protein levels is by modulating its transcription, thus indicating that regulation of its promoter plays a key role in its normal function. Indeed, CS patients with germline PTEN promoter mutations have a high prevalence of breast, thyroid and endometrial malignancies, and seem to have few of the other signs, thus, an understanding of the mechanism of PTEN dysfunction in these patients may lead to rational targeted therapies to treat or prevent malignancy. To this end, we performed several detailed studies of the PTEN promoter in which we demonstrated that Rosi(51),

Lovastatin, Simvastatin, Pravastatin and Fluvastatin(81) could induce PTEN

transcription; albeit through different mechanisms. In additional studies, we revealed a

novel method of CS progression due to mutations within the PTEN promoter that cause an inhibition of protein translation(9). Our data suggest that the agonists stated above, as well as a SREBP inducer, which can regulate PTEN’s transcription(81), would be highly effective within CS patients with a hemizygous deletion, a germline nucleotide alteration within the promoter, or in the circumstance where a PTEN mutation has yet to be

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identified. Additionally these therapeutic tools may also aid patients with sporadic tumors

with a somatic 5’-UTR VUS. However, despite these encouraging results, we must be

aware that these potential therapies may theoretically harm patients with germline

intragenic PTEN mutations or those with neoplasias by means of somatic intragenic mutations by raising levels of mutant, as well as WT protein. Additionally, our results indicate that one needs to be conscious that the combination of statins and a SREBP agonist may hinder the effect of each individual treatment.

Our data reiterate the importance of looking for variants within the PTEN

promoter and elsewhere in the 5’UTR within patients that have CS features yet do not

have a detectable mutation in its ORF. Furthermore, this knowledge can be extended to

other diseases and their respective susceptibility genes. Currently, promoters are rarely

analyzed in the clinical setting, therefore, it is very likely that nucleotide changes will be

identified in many other genes and these patients may also benefit from personalized

treatment given their promoter mutation status. The culmination of these works provides

more concrete evidence of the importance of the PTEN promoter in cancer development.

While these studies have resulted in some definitive answers to the role of the

PTEN promoter in CS development, they have also raised many interesting questions and

point the way to further research. In our first study, we tested several of the TZDs’ ability

to induce PTEN transcription and interestingly discovered that only one could induce

PTEN expression. It is common for agonists within a family of drugs to not signal exactly

the same, thus allowing them to express their individuality. In some cases this

individuality leads to detrimental side effects, which is in fact observed in the TZD

family. In 2000, Troglitazone (Rezulin) was pulled off of the market due to liver

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toxicity(82). Interestingly, to date, this has not been observed in the other TZDs.

Furthermore, a recent study demonstrated that Rosi (Avandia) increased an individual’s risk for heart complications(83), however, these results have yet to be replicated.

Furthermore, in this first meta-analysis, important studies which came to the opposite conclusions were not included in the metaanalysis. Understanding the mechanism behind these side effects, and untangling the differences between a family’s individual agonists may open the door to both new avenues of cancer treatment and more personalized health care. Moreover, this understanding will allow physicians to properly weigh the benefits against the known side effects prior to prescribing such a treatment.

Similar to the TZDs we observed the individuality of the statins; where all of the statins induced PTEN expression but Mevastatin. However, in contrast to the TZDs, only minor side effects have been observed with statin treatment, suggesting that the statins may be a safer clinical inducer of PTEN expression within CS patients. These agonists are currently being used clinically and are therefore readily available for patients who may greatly benefit from their use. Indeed, we did observe that normal control patient cell lines treated with the above statins increased PTEN expression (data not shown); however, we must acknowledge that previous studies have identified subsets of breast cancer patients that benefit from specific treatments based on tumor characteristics such as estrogen receptor (ER)(84) and p53(85) status. We studied statin signaling in both ER- positive (MCF-7 and T47D) and ER-negative (MDA-MB-435 and MDA-MB-231) cell lines. In both cell types, the statins could equally induce PTEN expression, suggesting that ER status is not relevant in statin signaling, which is in agreement with Mueck and colleagues’ results(86). Koyuturk et al further interrogated these results and demonstrated

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that neither ER or p53 status affected statin stimulation(87). Similarly, we observed that both statin and ALLN treatment in Saos-2 cells, which do not express p53, induced

PTEN expression (data not shown), thus suggesting that p53 is not necessary for PTEN induction by statins. These data suggest that a patient’s ER and p53 status does not affect statin upregulation of PTEN expression, an important observation given the commonality of somatic TP53 mutations in many sporadic malignancies.

The above analyses provide the necessary foundation for more in depth studies where future investigations will aim to further untangle the mechanisms behind PTEN promoter regulation. One of the most important studies to complete would be to determine what transcription factor(s) regulate PTEN between -893 and -755. Our results indicate that the protein or protein complex, which binds to this region is regulated by the statins, thus stressing its importance. As stated above, two transcription factors have been predicted to bind to this region, Sp1 and C-Myb; however, their relevance has not been determined. Several other transcription factors have been projected to bind to this region; however, they were only predicted by one software program indicating their importance may not be as significant. A few select predicted transcription factors are heat shock protein (HSP), heat shock factor (HSF), and retinoic acid receptor X (RXR). There are no data connecting these transcription factors with the statins or PTEN. We studied the connection between the PTEN promoter and HSP70; however, we were unable to demonstrate direct binding (data not shown). Moreover, our EMSA results suggest that

HSF2, and RXR are not the PTEN -893 to -755 transcription factors (data not shown). In order to concretely determine what transcription factor(s) bind to this region we plan to perform several biotinylated pull-down assays, followed by mass spectroscopy.

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Published research has demonstrated an association between statins and a few transcription factors(88, 89); however, the statins’ role in gene regulation has yet to be studied in great detail. We initially hypothesized that the statins signaled through SREBP to induce PTEN transcription, however our results indicated this to be false. Through further research, we were able to demonstrate that SREBP could independently induce

PTEN expression, thus suggesting a SREBP agonist may be clinically relevant to CS patients. Yet to date there is no known direct SREBP agonist, thus opening the door to allow the initiation of new drug discovery, which could be clinically relevant in a cancer setting. These data demonstrate that further studies need to be completed to determine the mechanism of PTEN transcriptional upregulation by the statins. Previous works have indirectly suggested that the statins may utilize the protein kinese A (PKA) pathway(90).

Additionally, PKA has been implicated in pathways upstream of PTEN transcription(91).

Indeed we observed that PKA inhibitors prevented the statins’ induction of PTEN expression when cells were treated simultaneously (data not shown). This suggests that the statins may be signaling down the PKA pathway to regulate PTEN expression, thus warranting further investigation.

Another area of interest is the mechanism of CS development in individual patients (Chapter 4). In that study, we analyzed a CS patient with a -764G/A nucleotide variant and speculated that their mechanism of CS development was due to inhibition of protein translation; however, further studies suggest that that may only be partially correct. Subsequent to the works demonstrated in Chapter 4, we performed an additional

EMSA to determine the significance of the -764G/A nucleotide variant. Nuclear protein was incubated with radiolabeled WT -893 to -755 PTEN promoter and either a WT or

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mutant -779 to -749 PTEN probe. As expected, the WT -893 to -755 PTEN promoter bound to the nuclear protein. Interestingly, the 30 base pair WT probe could not compete off the nuclear protein, however, the mutant probe did (Figure 5.1). This suggests that the mutation at -764G/A allowed for the addition of a novel transcription factor that does not normally bind to the PTEN promoter. Understanding the resulting outcome and the novel transcription factor bound to this region may allow us to manipulate normal signaling pathways and potentially produce more personalized treatment.

In conclusion, we have investigated several mechanisms of PTEN promoter

regulation and demonstrated novel modes of PTEN induction. We have shown that that

Rosi, Lovastatin, Simvastatin, Pravastatin and Fluvastatin can induce PTEN transcription.

Moreover, we also observed for the first time, SREBP’s ability to induce PTEN expression. Lastly, we demonstrated that PTEN 5’UTR mutations can cause an inhibition of protein translation and subsequently lead to CS progression. It is hoped that the culmination of these data will allow for both more personalized treatment options, as well as aid in better patient counseling. Furthermore, these studies also provide the groundwork for future experiments, such as those stated above, that will allow for novel molecular genetic-based therapies to become available.

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Figure 5.1 -764G/A PTEN Promoter Mutation Produces a Novel Transcription Factor

Binding Site

WT PTEN promoter from -893 to -755 was radiolabeled. Nuclear protein was either not incubated with the probe (Ng), or incubated with the probe (Bd) to test binding (lanes 1,

2). Competition assays were performed with -779 to -749 PTEN promoter that is either

WT (WT), or MT (MT) at the -764G/A position (lanes 3, 4). A representative blot from three individual experiments is displayed.

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