Pentoxifylline As an Adjuvant Treatment in Renal Cell Carcinoma

Pentoxifylline As An Adjuvant Treatment In Renal Cell Carcinoma

Item Type text; Electronic Dissertation

Authors Mastrandrea, Nicholas Joseph

Publisher The University of Arizona.

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Link to Item http://hdl.handle.net/10150/337293

PENTOXIFYLLINE AS AN ADJUVANT TREATMENT IN RENAL CELL CARCINOMA

Nicholas J. Mastrandrea

A Dissertation Submitted to the Faculty of the

DEPARTMENT OF PHARMACOLOGY AND TOXICOLOGY

In Partial Fulfillment of the Requirements

For the Degree of

DOCTOR OF PHILOSOPHY

In the Graduate College

THE UNIVERSITY OF ARIZONA

2014

THE UNIVERSITY OF ARIZONA GRADUATE COLLEGE

As members of the Dissertation Committee, we certify that we have read the dissertation prepared by Nicholas J Mastrandrea, titled Pentoxifylline as an Adjuvant Treatment in Renal Cell Carcinoma and recommend that it be accepted as fulfilling the dissertation requirement for the Degree of Doctor of Philosophy.

______Date: 9/25/14 Serrine S. Lau, Ph. D.

______Date: 9/25/14 Terrence J. Monks, Ph. D.

______Date: 9/25/14 Donna Zhang, Ph. D.

______Date: 9/25/14 Richard Vaillancourt, Ph. D.

______Date: 9/25/14 George Tsaprailis, Ph. D.

Final approval and acceptance of this dissertation is contingent upon the candidate’s submission of the final copies of the dissertation to the Graduate College.

I hereby certify that I have read this dissertation prepared under my direction and recommend that it be accepted as fulfilling the dissertation requirement.

______Date: 9/25/14 Dissertation Director: Serrine S. Lau

STATEMENT BY AUTHOR

This dissertation has been submitted in partial fulfillment of the requirements for an advanced degree at the University of Arizona and is deposited in the University Library to be made available to borrowers under rules of the Library.

Brief quotations from this dissertation are allowable without special permission, provided that an accurate acknowledgement of the source is made. Requests for permission for extended quotation from or reproduction of this manuscript in whole or in part may be granted by the copyright holder.

SIGNED: Nicholas J. Mastrandrea

ACKNOWLEDGEMENTS

First and foremost, I sincerely thank my mentor, Dr. Serrine Lau, for her guidance, patience and support throughout my graduate career. Dr. Lau provided me with enough freedom and funding to allow me to make this project my own. At times, she was the gas pedal in a stalled car. Other times, she was the brake pedal in a car out of control. Always, though, Dr. Lau put my training ahead of everything else. The independence and critical thinking I have gained here is a direct result of Dr. Lau's advising strategy; one that I am infinitely grateful for.

I would also like to thank my dissertation committee members, Dr. Terrence Monks, Dr. Donna Zhang, Dr. Richard Vaillancourt, and Dr. George Tsaprailis for their continuous advice and guidance throughout the years. They helped me to rethink about my data and suggested key experiments to perform in order to effectively demonstrate my hypothesis. Thank you for the time and effort each one of you have put into me and this project.

In addition, I am grateful for the numerous past and present members of the Lau and Monks labs for their training, friendship and support. Also, I would like to thank Dr. Nagle, Jaime Gard, David Chafin, and Ventana Medical Systems for our exciting, albeit challenging, collaboration. I would like to thank the staff at the shared cytometry, genomics, genetics, proteomics and animal cores here at the University of Arizona. Without your expertise and techniques, much of my data would not exist.

A special thanks goes to Wesley Cai who worked closely with me during the four years of his undergraduate career. It didn't take long to train Wesley, nor did it take long for him to demonstrate excellent scientific ability. Thus, I expected more from Wesley than is typical of an undergraduate and Wesley continually exceeded those expectations. More importantly, though, Wesley has been a great friend. I am very proud of him and his accomplishments and I look forward to what discoveries he will make in his career.

Finally, I would like to thank my family and friends for their support through these challenging six years. Thank you, Mom, Dad, and Gayle, for helping to form the man I am today. Thank you, Chris and Micah, Adrienne, Brad and Amanda, for your unconditional love and support. I couldn't have done this without the help of you all. To my friends, Patrick, Kuhlman, Pablo, Eric, Jon, Wnek and Wesley, thank you for all the shared experiences we had. I wish to never forget the memories we formed.

TABLE OF CONTENTS

LIST OF FIGURES AND TABLES ...... 10 LIST OF ABBREVIATIONS ...... 13 ABSTRACT ...... 17 CHAPTER 1:INTRODUCTION ...... 19 1.1 General Comments...... 19 1.2 Kidney Anatomy and Physiology ...... 20 1.3 Kidney Carcinogenesis ...... 24 1.3.1 von Hippel-Lindau Disease ...... 25 1.3.2 Tuberous Sclerosis Complex ...... 26 1.4 Cell Cycle and Its Regulation ...... 27 1.4.1 Restriction Point Regulation ...... 30 1.5 Cyclin D1 Regulation and Role in Cancer ...... 31 1.5.1 CCND1 Transcription via MAPK Signaling ...... 33 1.5.2 Cyclin D1 Translation via PI3K/AKT/mTOR/4EBP1 Signaling ...... 34 1.5.3 Cyclin D1 Nuclear Export and Proteasomal Degradation via GSK-3β Signaling ...... 37 1.6 The Eker Rat and Production of the QTRRE Cell Model ...... 41 1.6.1 Hydroquinone and Resulting Nephrotoxic Metabolites ...... 42 1.6.2 QTRRE Cell Model ...... 44 1.7 Human RCC Cell Models ...... 45 1.8 Anticancer Therapy ...... 46 1.8.1 Current FDA-Approved Targeted Therapies for RCC ...... 50 1.8.2 Sorafenib ...... 53 1.8.3 Current RCC Adjuvant Treatment Outlook...... 54 1.9 Pentoxifylline ...... 55 1.10 Dissertation Aims...... 61 CHAPTER 2: PENTOXIFYLLINE DECREASES CYCLIN D1, pRB LEVELS

AND ARRESTS RCC CELL MODELS IN THE G1 PHASE ...... 64 2.1 Introduction ...... 64 5

2.2 Materials and Methods ...... 67 2.2.1 Cell Culture...... 67 2.2.2 Western Blot Analysis ...... 67 2.2.3 QTRRE Expression Microarray Analysis ...... 68 2.2.4 RT-PCR ...... 69 2.2.5 Cell Cycle Analysis ...... 70 2.2.6 MTS Cell Viability Assay ...... 70 2.2.7 Statistics ...... 71 2.3 Results ...... 72 2.3.1 PTX, but not db-cAMP or theophylline, decreases cyclin D1 in QTRRE cells 72 2.3.2 PTX induces gene expression changes consistent with cell cycle arrest in QTRRE 74 2.3.3 PTX treatment for 24 hrs causes G1 phase arrest and cyclin D1 decrease in QTRRE, Caki-2 and ACHN but not in 786-O or HK-2 RCC cell models ...... 81 2.3.4 Cyclin D1 protein levels recover in 786-O, but not ACHN, at 24 hrs ...... 85 2.3.5 PTX effects on cell viability in ACHN ...... 87 2.3.6 PTX decreases cyclin D1 and pRB (Ser780) levels in ACHN in a time- and dose-dependent manner ...... 90 2.3.7 PTX induces G1-phase arrest in ACHN in a time- and dose-dependent manner 92 2.4 Discussion ...... 95 CHAPTER 3: PENTOXIFYLLINE DECREASES pAKT, pGSK-3β, p4EBP1, AND CYCLIN D1, WITHOUT AFFECTING CCND1 TRANSCRIPTION NOR CYCLIN D1 PROTEIN STABILITY ...... 100 3.1 Introduction ...... 100 3.2 Methods and Materials ...... 103 3.2.1 Cell Culture...... 103 3.2.2 RT-PCR ...... 103 3.2.3 Western Blot Analysis ...... 104 3.2.4 Plasmids and Site-Directed Mutagenesis ...... 105 3.2.5 Plasmid and siRNA Transfection ...... 106 3.2.6 Immunoprecipitation ...... 106 3.2.7 Statistics ...... 107

3.3 Results ...... 108 3.3.1 PTX has no effect on CCND1 transcription rates in QTRRE, ACHN or 786- O 108 3.3.2 PTX decreases pAKT (Thr308 and Ser473), p4EBP1 (Ser 65, Thr 70, and Thr 37/46) and cyclin D1 levels in a time- and dose- dependent manner in ACHN 110 3.3.3 PTX-mediated cyclin D1 decrease is independent of GSK-3β activity ...... 113 3.3.4 PTX-mediated cyclin D1 decrease is independent of DYRK 1B, and does not increase Thr 286 phosphorylation on cyclin D1 ...... 116 3.3.5 PTX-mediated cyclin D1 decrease is independent of Thr 286 phosphorylation 120 3.3.6 PTX-mediated cyclin D1 decrease is independent of proteasomal degradation and does not affect cyclin D1 stability ...... 123 3.4 Discussion ...... 127 CHAPTER 4: PENTOXIFYLLINE INCREASES THE EFFICACY OF SORAFENIB IN ACHN CELLS AND XENOGRAFTS IN MICE ...... 132 4.1 Introduction ...... 132 4.2 Materials and Methods ...... 134 4.2.1 Reagents...... 134 4.2.2 Cell Culture...... 134 4.2.3 MTS Cell Viability Assay ...... 134 4.2.4 Western Blot Analysis ...... 135 4.2.5 In Vivo Xenograft Study ...... 135 4.2.6 Xenograft Tissue Harvest and Analysis Preparation ...... 136 4.2.7 Immunohistochemistry ...... 137 4.2.8 Statistics ...... 138 4.3 Results ...... 139 4.3.1 Antiproliferative effects of sorafenib in ACHN are time- and dose- dependent 139 4.3.2 Combination of PTX + sorafenib treatment significantly decreases cyclin D1 and proliferation in ACHN cells ...... 141 4.3.3 Combination of PTX + sorafenib treatment inhibits ACHN tumor growth 145 4.3.4 Combination of PTX + sorafenib treatment decreases pRb, p4EBP1, and cyclin D1 levels in ACHN xenograft tissue ...... 148 4.4 Discussion ...... 151 CHAPTER 5: CONCLUDING REMARKS ...... 154 7

5.1 Antiproliferative effects of PTX in multiple RCC cancer models ...... 154 5.2 Targeting pAKT/p4EBP1/cyclin D1 in RCC ...... 156 5.3 PTX and combinatorial therapeutics in RCC...... 157 CHAPTER 6: FUTURE DIRECTIONS ...... 159 6.1 Overview ...... 159 6.2 PTX target identification...... 161 6.2.1 Introduction ...... 161 6.2.2 Methods ...... 162 6.2.3 Anticipated results ...... 163 6.3 Drug structure optimization ...... 165 6.3.1 Introduction ...... 165 6.3.2 Methods ...... 166 6.3.3 Anticipated Results ...... 167 6.4 Biomarker identification for predicting PTX response/sensitivity ...... 168 6.4.1 Introduction ...... 168 6.4.2 Methods ...... 168 6.5 PTX adjuvant activity alone and in combination with other targeted, anti-cancer therapeutics in ACHN and other cancer cell types ...... 170 6.5.1 Introduction ...... 170 6.5.2 Methods ...... 171 6.5.3 Anticipated results ...... 172 APPENDIX A: DIFFERENTIAL MODULATION OF p-4EBP1, p-ERK AND CYCLIN D1 BY AMUVATINIB/ERLOTINIB TREATMENT IN PTEN+ AND - PTEN PROSTATE CANCER CELLS ...... 173 A.1 Abstract ...... 173 A.2 Introduction ...... 175 A.3 Materials and Methods ...... 178 A.3.1 Cell Culture...... 178 A.3.2 Immunohistochemistry ...... 178 A.3.3 1D/2D Western Blot Analysis ...... 178 A.3.4 4EBP1 Immunoprecipitation ...... 180 A.3.5 Statistics ...... 180

A.4 Results ...... 181 A.4.1 LNCaP (PTEN -) display activated PI3K/AKT/mTOR signaling, whereas DU145 (PTEN +) display activated MAPK signaling ...... 181 A.4.2 4EBP1 expression is increased in prostate cancer compared to prostatic intraepithelial neoplasia (PIN)...... 181 A.4.3 Amuvatinib and combination amuvatinib + erlotinib treatment decreases p4EBP1 and cyclin D1 levels in LNCaP ...... 184 A.4.4 Erlotinib and combination erlotinib + amuvatinib treatment decreases pERK and cyclin D1 levels in DU145 ...... 186 A.4.5 Erlotinib and amuvatinib cause multiple phosphorylation loss of 4EBP1 .. 188 A.5 Discussion ...... 190

LIST OF FIGURES AND TABLES

Figure 1.1 Coronal section of the kidney and enlarged nephron, 23

Figure 1.2 Cell cycle regulation via the cyclins and cyclin dependent kinases, 29

Figure 1.3 Cyclin D1 activates CDK4 via heterodimerization, 32

Figure 1.4 Regulation of cyclin D1 synthesis, 36

Figure 1.5 Regulation of cyclin D1 nuclear transport and degradation, 40

Figure 1.6 Hydroquinone metabolism leads to formation and bioactivation of 2,3,5-tris-(glutathion-S-yl) hydroquinone (TGHQ), 43

Figure 1.7 Targeting the cell cycle in anti-neoplastic therapy, 48

Figure 1.8 Structural comparison between cAMP, PTX and PTX metabolite 1; lisofylline, 60

Figure 2.1 PTX, but not db-cAMP or theophylline, decreases cyclin D1 in QTRRE cells, 73

Table 2.1 Up- and downregulated gene cluster in QTRRE following 24 hr PTX treatment, 76

Table 2.2 Upregulated genes in QTRRE following 24 hr PTX treatment, 77

Table 2.3 Downregulated genes in QTRRE following 24 hr PTX treatment, 78

Figure 2.2 Downregulation of cell cycle related genes in QTRRE following 24 hr PTX treatment, 79

Figure 2.3 RT-PCR validation of select cell-cycle specific, downregulated genes in QTRRE following 24 hr PTX treatment, 80

Figure 2.4 Non-tumorigenic, HK-2, and RCC cell model cell cycle comparisons following PTX treatment, 83

Figure 2.5 Non-tumorigenic, HK-2, and RCC cell model comparisons following PTX treatment (1.0 mg/mL) for 24 hrs, 84

Figure 2.6 PTX effects on cyclin D1 in ACHN and 786-O, 86 10

Figure 2.7 Effect of PTX on mitochondrial dehydrogenase enzyme activity in ACHN cells, 89

Figure 2.8 PTX decreases cyclin D1 and pRB levels in ACHN, 91

Figure 2.9 PTX induces G1-phase arrest in ACHN in a time-dependent manner, 93

Figure 2.10 PTX induces G1-phase arrest in ACHN in a dose-dependent manner, 94

Figure 3.1 PTX yields no effect on CCND1 transcription rates, 109

Figure 3.2 PTX decreases pAKT and pGSK-3β levels in ACHN, 111

Figure 3.3 PTX decreases p-4EBP1 levels, concomitant with cyclin D1 decrease in ACHN, 112

Figure 3.4 GSK-3β inhibition via LiCl and does not affect PTX-mediated cyclin D1 decrease in ACHN, 114

Figure 3.5 GSK-3β knockout via siRNA does not affect PTX-mediated cyclin D1 decrease in ACHN, 115

Figure 3.6 Inhibition of DYRK1B via AZ191 does not affect PTX-mediated cyclin D1 decrease in ACHN, 118

Figure 3.7 PTX does not increase p-cyclin D1 Thr 286 levels following 2 hr and 4 hr doses, 119

Figure 3.8 Cyclin D1-HA mutant T286A decreases with PTX dose in ACHN, 122

Figure 3.9 MG-132 fails to completely rescue PTX mediated cyclin D1 decrease, 125

Figure 3.10 PTX has no effect on cyclin D1 stability, as demonstrated by T1/2, 126

Figure 4.1 Antiproliferative effects of sorafenib in ACHN cells, 140

Figure 4.2 Increased antiproliferative efficacy with PTX + sorafenib treatment in ACHN cells, 143

Figure 4.3 PTX pretreatment lowers the effective dose of sorafenib in ACHN cells, 144 11

Figure 4.4 In vivo ACHN tumor growth inhibition via PTX + sorafenib treatment, 146

Figure 4.5 PTX + sorafenib decrease pRB, p4EBP1, and cyclin D1 levels in ACHN xenograft tissue, via Western blot analysis, 149

Figure 4.6 PTX + sorafenib decrease cyclin D1 levels in ACHN xenograft tissue, via IHC analysis, 150

Figure A.1 PI3K/AKT/mTOR/4EBP1 and MAPK signaling differences in LNCaP and DU-145, 182

Figure A.2 4EBP1 levels in PIN and prostate cancer tissues, 183

Figure A.3 The effects of single and combination erlotinib (10 μM) and amuvatinib (10 μM) treatment in LNCaP, 185

Figure A.4 The effects of single and combination erlotinib (10 μM) and amuvatinib (10 μM) treatment in DU145, 187

Figure A.5 Combination erlotinib (10 μM) and amuvatinib (10 μM) treatment decreased 4EBP1 phosphorylation in LNCaP at 2.5 hr, 189

Figure A.6 Proposed signaling and drug perturbation schematic in DU- 145 (PTEN +) and LNCaP (PTEN -), 193

LIST OF ABBREVIATIONS

4EBP1 eukaryotic initiation factor 4E binding protein 1

AP-1 Activator Protein-1

AKT protein kinase B

APC anaphase promoting complex

ATM ataxia telangiectasia mutated

ATR ataxia telangiectasia mutated-related

AZ antizyme

BQ benzoquinone cAMP cyclic adenosine monophosphate

CCND1 cyclin D1 gene

CDK cyclin-dependent kinase

CRM1 exportin-1 db-cAMP dibutyryl-cyclic adenosine monophosphate

DMSO dimethyl sulfoxide

DOX doxorubicin

DYRK1B dual specificity tyrosine-phosphorylation-regulated kinase 1B

EGFR epidermal growth factor receptor eIF4E eukaryotic initiation factor 4E

EK eker rat germline mutation

ERK extracellular-signal-regulated kinase 1/2

ES enrichment score

FBS fetal bovine serum 13

FGFR(R) fibroblast growth factor (receptor)

FKHR forkhead in rhabdomyosarcoma

γ-GT gamma-glutamyl transpeptidase

G1 gap 1 of the cell cycle

G2 gap 2 of the cell cycle

GSH glutathione

GSK-3β glycogen synthase-3β

HA human influenza hemagglutinin

HIFα hypoxia-inducible factor α

HQ hydroquinone

IEF isoelectric focusing

IFN-α interferon-α

IP immunoprecipitation

IPG immobilized pH gradient

LOH loss of heterozygosity m/z mass-to-charge ratio

MALDI-TOF matrix-assisted laser desorption ionization-time of flight

MAPK mitogen activated protein kinase

MDR multidrug resistant mRCC metastatic renal cell carcinoma

MS mass spectrometry mTOR mammalian target of rapamycin

NSCLC non-small cell lung cancer 14

OSOM outer-stripe of the outer medulla

PAGE polyacrylamide gel electrophoresis

PBS phosphate-buffered saline

PDE phosphodiesterase

PDGF(R) platelet-derived growth factor (receptor)

PDK1 phosphoinositide-dependent kinase-1

PFS progression-free survival

PGP p-glycoprotein

PI3K phosphoinositide 3-kinase

PIN prostatic intraepithelial neoplasm

PIP3 phosphoinositol (3,4,5)-triphosphate

PTEN phosphatase and tensin homolog

PTM post-translational modification

PTX pentoxifylline

QTRRE quinol-thioether rat renal epithelial cell line

RB retinoblastoma

RCC renal cell carcinoma

RTK receptor tyrosine kinase

ROS reactive oxygen species

S-phase synthesis phase of the cell cycle

SCF skp, cullin, F-box containing complex

SFM serum-free media

TBS-T tris-buffered saline with 0.1% tween 15

TGF-α transforming growth factor α

TGHQ 2,3,5-tris-(glutathion-S-yl)hydroquinone

THEO theophylline

TIFP tumor interstitial fluid pressure

TSC tuberous sclerosis

UCC urothelial cell carcinoma

VCR vincristine

VEGF(R) vascular endothelial growth factor (receptor)

VHL von Hippel-Lindau disease

WCL whole cell lysate

ABSTRACT

Cyclin D1, a proto-oncogene, is required for progression from the G1 phase into the S phase of the cell cycle. Over-expression of cyclin D1 causes an increase in cell cycle progression and cell proliferation, implicating it in a variety of cancers including renal cell carcinoma (RCC). The rodent RCC cell model, QTRRE, and human RCC cell models, ACHN, 786-O and Caki-2, exhibit elevated levels of cyclin D1. Pentoxifylline

(PTX), a non-specific phosphodiesterase inhibitor, is an FDA-approved hemorheologic agent used to treat intermittent claudication, stemming from peripheral vascular diseases, as well as other diseases involving defective locoregional blood flow. Treatment of

QTRRE, ACHN, 786-O and Caki-2 with PTX caused a time- (0-24 hrs) and dose- (0-1.0 mg/mL) dependent decrease of cyclin D1 protein and p-Rb levels in whole cell lysate as well as cytosolic and nuclear fractions, albeit, to different extents within the models.

Concomitant with cyclin D1 and p-Rb decrease, enhanced G1 phase cell cycle arrest was observed in the RCC models.

Mechanistic studies in these RCC cell models were carried out to determine PTXs mechanism of action with regard to cyclin D1 protein level decrease. RT-PCR analysis showed no significant changes in cyclin D1 mRNA copy number in time- (0-24 hrs) and dose- (0-1.0 mg/mL) dependent PTX treatments. However, such treatments caused decrease in p-4EBP1 (Ser65), p-4EBP1 (Thr70), and p-4EBP1 (Thr37/46). Because

PTX’s ability to decrease cyclin D1 protein was prevented in the presence of the proteasome inhibitor, MG-132, studies were performed to determine whether cyclin D1 stability was decreased during PTX treatment. Cyclin D1 degradation is initiated by

phosphorylation of residue Thr286 by GSK-3β. Inhibition of GSK-3β with LiCl or knockdown via siRNA in the presence of PTX failed to block cyclin D1 decrease.

Moreover, PTX treatment in the presence of MG-132 revealed no significant increase in cyclin D1 p-Thr286 compared to control. Finally, using the protein synthesis inhibitor,

CHX, PTX caused no significant decrease in cyclin D1 t1/2 (wt-HA and T286A-HA) compared to control.

Sorafenib, a broad-spectrum (cRAF, bRAF, KIT, FLT-3, VEGFR-2, VEGFR-3, and PDGFR-β) kinase inhibitor, is FDA-approved for the treatment of RCC. Studies with sorafenib and PTX in the ACHN cell model were carried out to determine PTXs possible adjuvant role in inhibiting cell growth via cyclin D1 decrease and G1 phase arrest. MTS data showed PTX potentiates the anti-proliferative effects of sorafenib. PTX pre- treatment for 24 hrs was also lowered the effective dose of sorafenib from 50 µM to 5

µM. Further, ACHN xenograft tumor volumes from mice treated with PTX and sorafenib displayed significantly higher tumor growth inhibition compared to either drug treatment alone or vehicle. Finally, drug treated ACHN xenograft tissue displayed significantly lower cyclin D1, p-RB and p-4EBP1 levels. These results demonstrate a novel anticancer property of PTX and suggest its use as a possible adjuvant therapy in RCC treatment should be further explored.

CHAPTER 1:INTRODUCTION

1.1 General Comments

Cancer is the second leading cause of death in the United States with

585,720 estimated deaths and an estimated 1.67 million new cases in 2014

(cancer.gov 2014). Approximately 40.8% of American men and women will be diagnosed with some form of cancer at some point during their lifetime

(cancer.gov 2010). Cancer is a disease caused by an uncontrolled proliferation of abnormal cells. It can arise from a combination of environmental and genetic factors that cause mutations in various genes. These mutations lead to activation of oncogenes or deactivation of tumor suppressor genes, ultimately leading to disregulation of signaling pathways, uncontrolled cell growth and eventual tumor formation, invasion of surrounding tissue and metastasis.

Of the functional capabilities that cancers acquire, enhanced cell proliferation is one of the most extensively studied, and is the focus of this dissertation. Increased cell proliferation is achieved through various mechanisms but typically involve the disregulation of the cell cycle. This work investigates how a drug called pentoxifylline (PTX) interrupts the cell cycle in different cell and xenograft models of renal cell carcinoma (RCC), the most common form of kidney cancer, as well as explores its ability to synergize with sorafenib, an FDA- approved targeted treatment for RCC. Results from these studies suggest that

PTX may serve as an adjuvant treatment, alongside sorafenib and other targeted therapies, for the more effective treatment of this disease. 19

1.2 Kidney Anatomy and Physiology

The kidney is involved in a plethora of functions including the excretion of metabolic wastes, reabsorption of vital nutrients, regulation of extracellular fluid volume, regulation of blood pressure, electrolyte composition balance, acid-base homeostasis and the synthesis and release of various hormones (Klaassen

2001). The kidneys receive blood from the renal arteries and, despite their relatively small size, the kidneys receive approximately 20% of the cardiac output

(Walter 2004). The kidney is compartmentalized into three main sections including the cortex, medulla and papilla which receive 90%, 8% and 2% of the total blood flow to the kidney, respectively. The basic structural and functional unit of the kidney is the nephron (Figure 1.1), of which, there are approximately

800,000 - 1.5 million per human kidney (Guyton and Hall 2006). The beginning of the nephron is comprised of the renal corpuscle, made up of the glomerulus which is surrounded by the Bowman's capsule. Afferent arterioles supply blood flow to the glomerulus, where filtration of nutrients, wastes, electrolytes, water and small molecules first occur. The glomerular capillary wall serves as a barrier to the filtration of macromolecules, as well as regulated filtration of anionic molecules (Klaassen 2001). Filtrate then enters the renal tubule portion of the nephron for further processing.

The renal tubule consists of the proximal convoluted tubule, loop of Henle, and the distal convoluted tubule. The proximal convoluted tubule is divided into three segments, S1, S2, and S3. The S1 and S2 (pars convoluta) segments are

confined to the renal cortex and demonstrate higher cell complexity via their tall brush border membrane, highly developed vacuolar lysosomes, interdigitated basolateral membrane and densely packed mitochondria (Walter 2005; Klaassen

2001). The S3 segment (pars recta) descends into the outer stripe of the outer medulla (OSOM) and demonstrate lower cell complexity through decreased lysosomes and mitochondria, as compared to the pars convoluta. Despite this, the pars recta posses a highly developed brush border membrane (Walter 2005).

Following passage through the proximal convoluted tubule, filtrate travels through the loop of Henle which descends into the renal medulla via the thin descending limb, bends, and returns to the renal cortex via the thin and think ascending limbs. Water and ions are reabsorbed along the loop. Finally, the filtrate travels through the distal convoluted tubule where terminal reabsorption of nutrients and water occurs. The concentrated filtrate (urine) is passed through the collecting tubule and ureter, not considered parts of the nephron, to the urinary bladder for excretion (Hook and R. S. Goldstein 1993).

The proximal convoluted tubule is responsible for the majority of water and nutrient reabsorption. Roughly 66% of water and solutes are reabsorbed via transcellular transport, followed by active transport across the basolateral membrane via the Na+/K+/ATPase pump. In addition, nearly all organic solutes

(glucose, amino acids and citric acid cycle intermediates) and the majority of K+,

- - 3- 2+ 2+ HCO3 , Cl , PO4 , Ca , and Mg ions are reabsorbed in the proximal convoluted tubule (Klaassen 2001). Thus, disregulated homeostasis via injury to (acute tubular necrosis/acidosis) or tumor formation in the proximal convoluted tubule 21

will significantly impair water and solute balance as well as various other whole- kidney functions (Tomita, 2006). In addition, due to its high metabolic and transport capacity, and its ability to concentrate xenobiotics, the proximal convoluted tubule is most prone to toxicant-induced injury. Systemic glutathione

(GSH)-conjugated xenobiotics or metabolites are transported to the kidney in high concentrations for filtration from the blood and eventual excretion. Via the high gamma-glutamyltranspeptidase concentration in the S3 segment, the proximal convoluted tubule has the highest apical transport and accumulation of

GSH-conjugated molecules that may cause cytotoxicity, acute renal failure and possible tumorigenesis (Lau et al. 1988).

Figure 1.1 Coronal section of the kidney and enlarged nephron

The kidney is a highly compartmentalized organ, made up of three distinct regions: cortex, medulla and papilla. The structural and functional subunit of the kidney is the nephron, which spans the cortex and medulla regions of the kidney. Blood flow enters the nephron, via an afferent arteriole, through the glomerulus which is surrounded by the Bowman's capsule. Filtrate then passes, in the direction of the arrows, through the proximal convoluted tubule, followed by the loop of Henle and into the distal convoluted tubule. Concentrated filtrate (urine) finally flows through the collecting tubule towards the ureter and is stored in the urinary bladder for excretion. Water and nutrient reabsorption occur throughout the nephron. 23

1.3 Kidney Carcinogenesis

In the United States, 64,000 new cases of kidney cancer are estimated in

2014, and 14,000 deaths will occur from the disease (cancer.gov 2014). It is the seventh most common cancer in men and the ninth most common in women. In the last decade, new kidney cancer cases have been rising on average 1.6% each year (cancer.gov 2014). The most common types of kidney cancer are renal cell carcinoma (RCC) and urothelial cell carcinoma (UCC). RCC represents approximately 90-95% of all primary renal cancers and arises in the proximal convoluted tubule (Curti, 2004). UCC accounts for the majority of the remainder of kidney cancer cases and arises in the renal pelvis, a part of the ureter. Less common types of kidney cancer include squamous cell carcinoma, Bellini duct carcinoma, mesoblastic nephroma and mixed epithelial stromal tumor.

RCC is considered one of the deadliest of cancers affecting the genitourinary tract (Ramana, 2012). Despite the earlier detection of smaller kidney tumors, the rate of RCC-related mortality has increased, suggesting that recurrence and advanced disease are responsible for mortality (Hollingsworth et al., 2006; Rini et al., 2009). Patients with metastatic RCC (mRCC) have a 5-year survival rate of less than 10%. RCC occurs with twice the frequency in men than in women and studies have shown that women have a less severe prognosis and higher rates of survival following RCC recurrence compared to men (Jemal et al.

2004.; Onishi et al. 2002). The main risk factors for RCC are smoking, obesity and hypertension (Häggström et al., 2013).

1.3.1 von Hippel-Lindau Disease In addition to environmental factors, some genetic factors have been implicated in the formation of RCC. Congenital diseases including von Hippel-

Lindau disease (VHL) and autosomal dominant polycystic disease are associated with human cases of RCC. 80% of human RCC cases reveal a mutation in the

VHL tumor suppressor gene, most commonly arising from a deletion in chromosome 3 p (Gnarra et al., 1996). The VHL gene locus undergoes loss of heterozygosity (LOH) early in human RCC and its tumorigenic inactivation is highly specific to the kidney (Walker 1998). The protein encoded by VHL is an E3 ubiquitin ligase (pVHL) responsible for the degradation of hypoxia-inducible factor α subunits (HIFα), a transcription factor that plays a central role in oxygen homeostasis (Maxwell et al. 2001). In addition, pVHL regulates cell cycle arrest though stabilization of the cyclin-dependent kinase inhibitor, p27 (Pause et al.,

1998). Vascular endothelial growth factor (VEGF), a downstream transcriptional target of HIFα, is the responsible for angiogenesis (Siemeister et al., 1996).

Thus, loss of functional pVHL leads to enhanced cell cycle progression and angiogenesis, two requirements for the formation of a tumor. The human renal cell carcinoma cell models used in this work are VHL null and are further covered in 1.7. Contrary to human RCC, the VHL tumor suppressor gene is not a primary target in rodent renal tumor formation (Kikuchi et al., 1995; Walker et al., 1996).

VHL mutations in the rat are rare (Shiao et al., 1997) and VHL knockout mice do not develop spontaneous RCC more than their wild-type littermates (Gnarra et al., 1997; Haase et al., 2001).

1.3.2 Tuberous Sclerosis Complex Tuberous sclerosis complex (TSC) is an autosomal dominant hereditary disease characterized by spontaneous mutation in the TSC-1 gene (human

9q34), encoding the protein hamartin, and the TSC-2 gene (human 16p13), encoding the protein tuberin (Kida et al., 2005). These proteins form a tumor suppressing complex that inhibit the mammalian target of rapamycin (mTOR), a signaling protein implicated in tumor development, and drug resistance against chemotherapy and radiation therapy (Narayanan, 2003). TSC manifests as cystic lesions in the brain, heart, lungs, skin and kidney (Gomez, 1991), and LOH at either the TSC-1 or TSC-2 gene locus have been identified in preneoplastic lesions of TSC patients (Carbonara et al., 1994; Green et al., 1994; Henske et al., 1996). 2% to 4% of patients with TSC develop RCC, higher than the estimated incidence in the general population (Ljungberg et al., 2011; Yang et al.,

2014). In the rodent, TSC-2 is a major target for both spontaneous (in rats and mice) and chemically induced (rats only) RCC formation (Kobayashi et al., 1999,

1995; Onda et al., 1999; Urakami et al., 1997; Yeung et al., 1994). The rodent

RCC cell model used in this work is TSC-2 null, further covered in 1.6.2.

1.4 Cell Cycle and Its Regulation

Most, if not all, cancers are believed to acquire the same set of functional capabilities including: self-sufficiency in growth signals and limitless replicative potential, insensitivity to anti-growth signals and evasion of apoptosis, sustained angiogenesis, and tissue invasion and metastasis (Hanahan and Weinberg,

2000). Self-sufficiency in growth signals results in an enhanced cell proliferation rate and is one of the most studied aspects of cancer research. Increased cell proliferation rate is achieved through a multitude of mechanisms but typically involves the disregulation of the cell cycle.

The cell cycle can be divided into two main periods: interphase, where the cell prepares for division, and mitosis, where division of a cell produces two daughter cells. Interphase can be broken down into 3 distinct periods: gap 1

(G1), where the cell grows in size and synthesizes mRNA and proteins in preparation for subsequent phases, synthesis phase (S phase), where DNA undergoes replication, and gap 2 (G2), where the cell readies itself for mitosis

(Figure 1.2A). Between these phases exist important checkpoints to ensure proper actions have occurred before continuing into the next subsequent phase.

The G1 checkpoint (also known as the restriction point; detailed in 1.4.1) controls

G1/S transition and ensures the cell is properly readied for DNA synthesis.

Beyond this point, the cell is committed to carrying out the next phases of the cell cycle and mitogenic signals are no longer required (Pardee, 1989). The G2 checkpoint ensures that DNA was successfully replicated and is largely free of

damage, in preparation for mitosis (Gould and Nurse, 1989). Finally, the metaphase checkpoint (or mitotic spindle checkpoint), occurs within mitosis and ensures the chromosomes are properly aligned at the mitotic plate, in preparation for the even division of chromosomes between the two daughter cells (Ciosk et al., 1998). As cells exit mitosis, the cell cycle is resets, allowing the establishment of a new, competent replication state in G1 phase.

The cell cycle is regulated by a concert of proteins, including cyclins (A-,

B-, D-, and E-type), cyclin-dependent kinases (CDK1, CDK2, CDK4, and CDK6), and cyclin dependent kinase inhibitors [CDKI; Cip/Kips (p21 and p27) and Ink4s

(p16)] (Malumbres and Barbacid, 2009). CDK and CDKI levels remain largely constant throughout the cell cycle, however, relative cyclin protein levels are highly dynamic (Figure 1.2B) (Malumbres and Barbacid, 2009). Activation of the

CDKs is dependent on binding the cyclin protein co-factors and, thus, proper cell cycle regulation is achieved when the respective cyclins are not only synthesized, but also degraded, at the proper times (Morgan, 1995). The E3- ubiquitin ligase complexes Skp1, Cullin and Fbox (SCF), and the Anaphase-

Promoting Complex (APC) are largely responsible for governing the stability of various cell cycle related proteins (van Leuken et al., 2008).

Figure 1.2 Cell cycle regulation via the cyclins and cyclin dependent kinases A concert of cyclin dependent kinases (CDKs) and their respective cyclin protein co-factors govern transition through the cell cycle (A). Following mitosis, the cell enters Gap1 (G1) phase where it begins to grow and prepare for division. CDK4/6-cyclin D (light blue) initiate progression into the synthesis (S) phase. G1 to S phase transition is completed by CDK2-cyclin E (black). CDK2-cyclin A (dark blue) carry the cell through S phase. CDK1-cyclin B (yellow) control S phase to Gap2 (G2) transition and entry into mitosis. Throughout the cell cycle, CDK levels remain constant, however, relative cyclin expression levels change with respect to cell cycle stage (B). Proper cell cycle regulation requires tight control over the respective cyclin expression levels.

1.4.1 Restriction Point Regulation Considered the rate-limiting step of the cell cycle, the restriction point allows for G1 to S-phase transition. Once a daughter cell reaches adequate size, and if mitogens are present, cyclin D1 is synthesized. Cyclin D1 complexes with and activates CDK4/6 in the cytosol (Figures 1.3B). Activated cyclin D1-CDK4/6 then enters the nucleus where it phosphorylates the tumor suppressor retinoblastoma (Rb) at residue Ser 780. Unphosphorylated Rb blocks the G1/S- phase transition by binding the E2F transcription factor, thereby inhibiting it (Luo et al., 1998; Zheng and Lee, 2001). Phosphorylation of Rb by cyclin D1-CDK4/6 causes dissociation between Rb and E2F, allowing for the E2F-mediated transcription of genes that promote G1/S-phase transition (e.g. cyclin E) (Harbour and Dean, 2000). Cyclin E binds and activates CDK2. The activated cyclin E-

CDK2 complex further phosphorylates Rb, leading to a hyperphosphorylated state, fully inhibiting E2F and delivering the cell into the S-phase. Further, cyclin

E-CDK2 phosphorylates the CDK inhibitor, p27, leading to degradation of p27

(Sheaff et al., 1997). In addition to cyclin E, other E2F genes are transcribed whose products aid in cell cycle progression into and in the S-phase, namely those involved in nucleotide metabolism and DNA synthesis, including DNA polymerase, thymidine kinase, dihydrofolate reductase and cdc6) (Nevins, 1998).

1.5 Cyclin D1 Regulation and Role in Cancer

Because of its important role in restriction point regulation, cyclin D1 is a tightly regulated protein with a half-life of approximately 20-60 min, depending on cell type (Dal Col and Dolcetti, 2008; Koepp et al., 1999). Cyclin D1 contains three important motifs and two important residues that govern its expression levels, cytosolic/nuclear localization, binding partner affinity and role in activating

CDK4/6 (Figure 1.3A). Cyclin D1 binds Rb near the N-terminus via the LXCXE motif and helps to facilitate cyclin d1-CDK4/6 phosphorylation. Binding to CDK4/6 is mediated via the cyclin box. Mutation of residue K112 within this domain to

E112 inhibits cyclin D1s ability to activate CDK4/6 (Landis et al., 2006; Li et al.,

2006). The cyclin box is also necessary for binding of the CDK inhibitors, p21 and p27. Near the C-terminus occurs the PEST domain that is rich in proline, glutamate, serine and threonine, characteristic of proteins that are rapidly turned over. Within the PEST domain is T286, a residue that is important in the nuclear export and proteasomal degradation of cyclin D1 (detailed in 1.5.4). The PEST domain is also important in binding of p21 and p27.

The deregulation of cyclin D1 expression can directly lead to some of the hallmarks of cancer (Hanahan and Weinberg, 2000). Overexpression of cyclin D1 can be attributed to many factors including increased transcription, translation, and protein stability, covered in detail below.

Figure 1.3 Cyclin D1 activates CDK4 via heterodimerization A. Cyclin D1 schematic displaying conserved motifs and locations of binding partners. Cyclin D1 binds Rb near the N-terminus via the LXCXE motif. The cyclin box mediates CDK4/6 binding and is necessary for interaction with the CDK inhibitors p21 and p27. K112 is required for CDK4/6 activation as K112E mutant is able to bind, but not activate CDK4/6. The C-terminal PEST domain is rich in proline, glutamate, serine and threonine which is characteristic of proteins that are rapidly turned over. Phosphorylation at T286 by GSK-3β triggers nuclear export via CRM1 and subsequent ubiquitin-mediated proteasomal degradation. T286A mutant confers higher stability than wt and is constitutively located in the nucleus. B. Ribbon diagram of the CDK4 (cyan)/cyclin D1 (orange) heterodimer. The N- and C-terminal lobes of the kinase are labelled as are key secondary structural elements (Day et al., 2009).

1.5.1 CCND1 Transcription via MAPK Signaling CCND1, the gene encoding cyclin D1, is a well established human oncogene and has a high frequency of amplification and overexpression in a wide variety of cancers including breast, lung, melanoma, bladder, esophageal, colorectal, and mantle cell lymphoma, among others (Beroukhim et al., 2010;

Bignell et al., 2010; Curtin et al., 2005; Santarius et al., 2010). The mitogen- activated protein kinase (MAPK) pathway regulates a plethora of cellular activities including gene expression, metabolism, and cytoskeletal changes, implicating it in cell proliferation, survival, differentiation and senescence.

Activation of this pathway occurs via receptor tyrosine kinases (e.g. epidermal growth factor; EGFR), which become activated when growth factors and other mitogenic ligands bind. Upon receptor activation, the Ras/Raf/MEK/ERK MAPK cascade of downstream enzymes become activated, sequentially. ERK activation ultimately leads to activation of various transcription factors including myc and

AP-1 that control a plethora of transcriptional targets, including CCND1 (Figure

1.4). Members of the MAPK signaling pathway (Ras, B-Raf) are found mutated or overexpressed in a multitude of human cancers and represent common targets for treating cancer. Hyperactivation of the MAPK pathway is directly correlated with overexpression of cyclin D1 protein levels and enhanced cell-cycle progression (Dhillon et al., 2007).

1.5.2 Cyclin D1 Translation via PI3K/AKT/mTOR/4EBP1 Signaling Cyclin D1 protein levels rely not only on CCND1 transcription but also mRNA translation. The PI3K/AKT/mTOR pathway regulates cell growth, proliferation, differentiation, motility and survival. Parallel to the MAPK pathway, the PI3K pathway is also found overexpressed in various human cancers. In addition to upstream receptor activation, loss of the TSC and PTEN tumor suppressors also contribute to increased pathway activation and occur in a number of cancers, including RCC (Hager et al., 2007). Downstream of the

PI3K/AKT/mTOR pathway, eukaryotic initiation factor 4E (eIF4E) binding protein

1 (4EBP1) regulates cyclin D1 expression through binding eIF4E and inhibiting eIF4E-mediated 5' cap-dependent translation (De Benedetti and Graff, 2004;

Sonenberg and Gingras, 1998). Hyperphosphorylation of 4EBP1 (at residues Ser

65, Thr 70, and possibly others), via upstream PI3K/AKT/mTOR signaling activation, causes decreased affinity between 4EBP1 and eIF4E, allowing for translation of cyclin D1 to occur (Figure 1.4) (Faivre et al., 2006). 4EBP1 expression was found to be associated with poor prognosis in several human tumors, such as breast, colon, ovarian, prostate, and renal cancers (Armengol et al., 2007; Coleman et al., 2009; No et al., 2011). The phosphorylation of 4EBP1 was also found to be associated with chemoresistance in various cancer types

(No et al., 2011).

Increased PI3K/AKT/mTOR signaling plays a significant role in the tumorigenesis of RCC and represents a set of amenable proteins in the therapeutic targeting of RCC (Cho, n.d.; Conti et al., 2013; Habib and Liang, 34

2014). Of note, two of the seven FDA approved targeted therapies for advanced and metastatic RCC, inhibit mTOR, effectively reducing 4EBP1 phosphorylation and subsequent translation of proteins, like cyclin D1, believed to contribute to cancer progression (covered in 1.8.1) (Faivre et al., 2006; Jastrzebski et al.,

2007).

Figure 1.4 Regulation of cyclin D1 synthesis

MAPK activation of ERK leads to transcriptional activation of the CCND1 gene, often found amplified and overexpressed in a multitude of human cancers. In the cytosol, cyclin D1 synthesis is regulated by eIF4E-mediated 5' cap-dependent translation of cyclin D1 mRNA. eIF4E is inhibited by 4EBP1. Activation of the PI3K/AKT/mTOR pathway leads to hyperphosphorylation of 4EBP1, reducing its affinity for eIF4E. This, in turn, increases the synthesis of cyclin D1 protein levels.

1.5.3 Cyclin D1 Nuclear Export and Proteasomal Degradation via GSK-3β Signaling

Control over cyclin D1 protein stability is one of the most crucial aspects of its regulation. With a half-life of approximately 30 - 45 min, cyclin D1 degradation is tightly controlled (Diehl et al., 1998, 1997). Inactivation of cyclin D1 via nuclear export and subsequent 26S proteasomal degradation is controlled by glycogen synthase kinase 3β (GSK-3β) (Diehl et al., 1998). In response to DNA damage or nutrient depravation, GSK-3β phosphorylates cyclin D1 at Thr286 in the nucleus

(Guo et al., 2005). This phosphorylation event unearths a nuclear export signal recognized by the nuclear exportin, CRM1, which transports cyclin D1 out of the nucleus (Alt et al., 2000; Benzeno and Diehl, 2004). This, in turn, enables the cytosolic E3 ubiquitin ligase, SCFFBX4/αβ-crystallin to bind to and ubiquitinate phosphorylated cyclin D1, targeting it for 26S proteasomal degradation (Figure

1.5) (Barbash and Diehl, 2008; Diehl et al., 1997; Lin et al., 2006). Several proteins, including AKT, regulate the activity of GSK-3β through Ser9 phosphorylation, an auto-inhibitory residue (Stambolic and Woodgett, 1994;

Zhang et al., 2003).

Disregulation of cyclin D1 nuclear export increases its oncogenic potential

(Benzeno et al., 2006), suggesting that nuclear retention of cyclin D1, resulting from altered nuclear trafficking and proteolysis, is critical for the manifestation of its oncogenicity (Kim and Diehl, 2009). Research using a mutated form of cyclin

D1 at Thr286 to Ala (D1-T286A), which cannot be phosphorylated by GSK-3β,

effectively demonstrate that CRM1 nuclear export and SCFFBX4/αβ-crystallin- mediated ubiquitination are crucial in limiting cyclin D1s oncogenic potential. D1-

T286A remained nuclear throughout the cell cycle and overexpression of the nuclear cyclin D1-T286A in murine fibroblasts resulted in cellular transformation and promoted tumor growth in immune compromised mice (Alt et al., 2000). Further, transgenic mice that overexpress D1-T286A developed mammary adenocarcinoma with a shorter latency relative to mice overexpressing the wild-type cyclin D1 (Lin et al., 2008). These observations demonstrate that subcellular localization and stabilization of cyclin D1 through its altered nuclear trafficking and proteolysis exert profound effects on tumorigenesis. Indeed, several mutations that impair phosphorylation-dependent nuclear export of cyclin

D1 have been identified in endometrial cancer (Ikeda et al., 2013; Moreno-Bueno et al., 2003) as well as in esophageal carcinoma and esophageal cancer-derived cell lines (Benzeno et al., 2006).

Among more than 100 polymorphisms identified in cyclin D1, the G870A mutation has gained the most attention due to its contribution to cyclin D1 alternative splicing, resulting in a truncated cyclin D1 (cyclin D1b) protein

(Betticher et al., 1995). Cyclin D1b can form an active complex with Cdk4 (Lu et al., 2003; Solomon et al., 2003) but lacks the C-terminal Thr286 residue whose phosphorylation is critical for nuclear export and subsequent cytoplasmic degradation of cyclin D1. Similar to cyclin D1 T286A, this structural difference of cyclin D1b renders the protein refractory to nuclear export through the cell cycle, which increases its oncogenic potency (Alt et al., 2000; Lu et al., 2003). High 38

levels of cyclin D1b have been reported to be associated with a poor prognosis for breast cancer patients (Abramson et al., 2010; Asiedu et al., 2014; Taneja et al., 2010; Zhu et al., 2010) as well as non-small cell lung cancer patients (Li et al., 2008). Administration of cyclin D1b siRNA inhibited breast tumor growth in nude mice and cyclin D1b siRNA synergistically enhanced the cell killing effects of doxorubicin in cell culture, with this combination also significantly suppressing tumor growth in the mouse model (Wei et al., 2011).

Figure 1.5 Regulation of cyclin D1 nuclear transport and degradation

Cyclin D1 lacks a nuclear localization signal, and, thus, must bind CDK4/6, and possibly other proteins, for transport into the nucleus. In the nucleus, activated cyclin D1/CDK4/6 complex inhibits Rb via phosphorylation at Ser780. This leads to decreased binding and sequestration of the E2F transcription factor by Rb, ultimately leading to transcription of E2F downstream genes (e.g. cyclin E) that serve to progress the cell through G1 to S phase transition. GSK-3β phosphorylates cyclin D1 at Thr286, causing nuclear export through CRM1. Phosphorylated cyclin D1 is ubiquitylated in the cytosol by the E3 ubiquitin ligase, FBX4, targeting it for 26S proteasomal degradation (Alao, 2007).

1.6 The Eker Rat and Production of the QTRRE Cell Model

The Eker rat (TSC-2EK/+) is a derivative of the Long Evans strain, bearing a

6.3 kb intracisternal-A partial insertion containing multiple stop codons that interfere with proper TSC-2 transcription (Eker et al. 1981; Kobayashi et al.

1995). Compared to wild-type rats, which display spontaneous RCC tumor formation in less than 0.05%, nearly all TSC-2EK/+ rats develop RCC by 12 months of age (Eker et al. 1981; Yeung et al. 1995). Thus, the inactive allele of

TSC-2 gene makes the Eker rat a useful model in studying the pathology and signal transduction involved in renal tumor formation. Further, TSC-2EK/+ rats display higher predisposition to spleen, pituitary gland, and uterine tumor development (Kobayashi et al. 1995; Yeung et al. 1994; Yeung et al. 1995).

Homozygous Eker mutation (TSC-2EK/EK) is embryonic lethal by day 9.5 - 13.5, demonstrating the important role of tuberin in proper development (Rennebeck et al., 1998). On a cellular, molecular, and phenotypic scale, renal tumors derived from TSC-2EK/+ rats are significantly similar to that of human RCC (Everitt et al.

1992; Everitt et al. 1995; Yoon et al. 2002). RCCs from both the TSC-2EK/+ rat and human derive from epithelial cells that line the proximal convoluted tubule of the nephron, overexpress transforming growth factor alpha (TGF-α) and cyclin

D1, and lack Ras and P53 mutations (Hard, 1986; Kobayashi et al., 1999; Recio et al., 1991; Walker et al., 1991).

1.6.1 Hydroquinone and Resulting Nephrotoxic Metabolites Hydroquinone (HQ; 1,4-benzenediol), the simplest of quinones, is found ubiquitously in nature, as well in the chemical industry. Roughly 35,000 tons of

HQ are produced annually (Westerhof and Kooyers, 2005). Some of its industrial uses include food preservation, rubber manufacturing, skin-care product depigmentation, photograph development, and it is a component of unfiltered cigarette smoke (Monks et al., 1992; Pryor, 1997). HQ can be oxidized by cytochrome P450 (1A1, 2E1, and 3A4), prostaglandin H synthase, or myeloperoxidase enzymes to 1,4-benzoquinone (BQ) (Figure 1.7); which can chemically react with glutathione (GSH) to form 2-(glutathion-S-yl)HQ and a multitude of substituted GSH adducts (Hill et al. 1993; Lau and Monks 1987; Lau et al. 1988; Monks et al. 1992; Subrahmanyam et al. 1990). Additional GSH conjugation leads to formation of 2,3,5-tris-(glutathion-S-yl)hydroquinone (TGQH)

(Figure 1.7), the most potent nephrotoxic adduct (Lau et al., 1988a). TGHQ is further metabolized via the mercapturic acid pathway in the OSOM to produce highly reactive HQ-cysteine conjugates, capable of redox cycling, resulting in reactive oxygen species (ROS) formation, as well as macromolecular adduction

(Dekant, 1996; Duffel and Jakoby, 1982; Monks and Lau, 1998, 1997, 1992).

Thus, GSH-HQ conjugates are selective nephrotoxicants, particularly to the epithelial cells lining the proximal convoluted tubule in the OSOM, and demonstrate significant tumorigenicity (Whysner et al., 1995).

Figure 1.6 Hydroquinone metabolism leads to formation and bioactivation of 2,3,5-tris-(glutathion-S-yl) hydroquinone (TGHQ) Hydroquinone is conjugated to GSH in liver. GSH conjugates are targeted to the kidney for clearance. GSH-HQ is further metabolized by γ-glutamyl transpeptidase (γ-GT) and cysteinylglycine dipeptidase enzymes in the S3 segment of the OSOM. HQ-cysteine is recognized by amino acid transporters in proximal tubules in the OSOM. Following re-uptake, the highly reactive HQ- cysteine conjugates are capable of redox cycling, resulting in the formation of reactive oxygen species (ROS), as well as conjugate adduction of macromolecules (Cohen 2009).

1.6.2 QTRRE Cell Model Previously, we have demonstrated TGHQ to be mutagenic, causing transformation of primary renal epithelial cells isolated from the TSC-2EK/+ rat

(Yoon et al. 2001). Immortalization of TSC-2EK/+ rat primary proximal convoluted tubule cells following transformation with TGHQ established the quinol-thioether rat renal epithelial (QTRRE) cell model (Yoon et al. 2001). QTRRE cells posses cytogenetic alterations, LOH at the TSC-2 gene locus (resulting in TSC-2Ek/-), and are null for tuberin expression (Yoon et al. 2001). Further, QTRRE cells are tumorigenic in athymic nude mice and produce tumors that resemble RCC cells

(Patel et al., 2003). TGHQ administration to Tsc-2EK/+ rats induces a sustained regenerative hyperplasia at sites that subsequently give rise to tumors (Lau et al.

2001; Yoon et al. 2001; Yoon et al. 2002). Similar to the QTRRE cells, preneoplastic lesions and tumors formed in TGHQ- treated TSC-2EK/+ rats displayed a LOH of the TSC-2 wild-type allele, subsequent loss of tuberin protein expression, high ERK kinase activity and high cyclin D1 expression (Lau et al.

2001; Yoon et al. 2002). In addition to inhibiting mTOR signaling, mechanistic studies in the QTRRE cells revealed tuberin to also be a suppressor of

Raf/MEK/ERK MAPK signaling (Yoon et al. 2004). QTRRE cells express constitutively high levels of activated ERK and B-Raf, as well as cyclin D1 and transient transfection of TSC-2 cDNA resulted in significant decrease of ERK and

B-Raf activity, concomitant with a decrease in cyclin D1 protein levels (Yoon et al. 2004).

1.7 Human RCC Cell Models

Compared to the rodent QTRRE cell model null for TCS-2, the ACHN human RCC cell model most used in these studies are null for VHL, reflective of

RCC tumors found in humans (Gnarra et al. 1996; Walker et al. 1998). The

ACHN RCC cell line was isolated via malignant pleural effusion of the lung in a

22-year-old Caucasian male with widely metastatic renal cell carcinoma (autopsy confirmed). The Caki-2 and 786-O RCC cell lines (VHL positive) were isolated from the primary tumor of a 69 year old and 58 year old Caucasian male, respectively. The ACHN, Caki-2 and 786-O cell lines are among some of the most commonly used as human RCC models.

Human VHL RCC and rodent Tsc-2 RCC not only share clear cell pathology, but also a number of common downstream targets. VHL and Tsc-2 null RCC models both result in accumulation of HIFα leading to increased protein levels of VEGF, as well as activation of MAPK and PI3K/AKT/mTOR/4EBP1 signaling, leading to overexpression of cyclin D1 and enhanced cell proliferation rates (Brugarolas et al., 2003; Liu et al., 2003; Rebuzzi et al., 2007). In addition, like QTRRE, ACHN, Caki-2 and 786-O form subcutaneous tumors in athymic mice. These striking similarities between human VHL RCC and our rodent Tsc-2

RCC model allow for plausible extrapolation between the two models.

1.8 Anticancer Therapy

Because cancer is a manifestation of cell cycle disregulation, therapies often interfere with one or multiple processes involved in cell cycle progression

(Figure 1.6). Traditional chemotherapies represent a majority of current cancer treatments. Alkylating agents (nitrosureas, nitrogen mustards) alter DNA and block replication. Anti-metabolites (hydroxyurea, 5-fluorouracil) substitute precursors of nucleic acid synthesis and also block DNA replication.

Topoisomerase I inhibitors (camptothecin) and II inhibitors (doxorubicin, mitoxantron) interfere with the ligation step of DNA synthesis and eventually cause apoptosis and cell death. Tubulin polymerization inhibitors (taxol) and tubulin depolarization inhibitors (vincristine, vinblastine) interfere with correct mitotic spindle formation, inhibiting mitosis. These chemotherapies yield significant adverse side-effects, notably immunosupression, and confer little to no specificity to tumor cells compared to normal cells.

More recent molecules have been developed to inhibit specific regulators of cell cycle and cell death, including kinases (MAPKs, PI3K, RTKs, etc) and phosphatases. Over 250 kinase inhibitors (both small molecule and antibody- based) are currently in clinical trials (Akritopoulou-Zanze and Hajduk, 2009;

Eglen and Reisine, 2011). About 23 muti-kinase-targeted inhibitors have reached the market (some covered in 1.8.1). CDK-specific inhibitors are newer and less developed, compared to other multi-kinase inhibitors. These drugs seek to inhibit the very class of proteins responsible for cell cycle progression and are currently

being investigated in both pre- and clinical settings (Abate et al., 2013; Diaz-

Moralli et al., 2013; Esposito et al., 2013; Gallorini et al., 2012). Initially, pan-CDK inhibitors failed to meet expectations, however, a second generation of highly specific cyclin D1/CDK4/6 inhibitors are proving to be more efficacious in slowing the growth of tumors (Casimiro et al., 2014). For example, a phase II study showed that PD0332991; PalbociclibTM (Pfizer), a selective CDK4/6 inhibitor, doubled progression free survival in patients with advanced breast cancer when combined with the aromatase inhibitor, letrozole; FemaraTM (Novartis), compared to letrozole alone (20 months vs 10 months) (Finn et al., 2012). Interestingly,

PD0332991 also demonstrated G1 cell cycle arrest, induction of late apoptosis, and blockade of RB phosphorylation in numerous RCC cell lines (Logan et al.,

2013). The ultimate goal of these targeted therapies (either multi-kinase or CDK specific) is to increase cancer-cell specificity; ultimately increasing efficacy as well as decreasing adverse effects.

Figure 1.7 Targeting the cell cycle in anti-neoplastic therapy

Various small molecule inhibitors have been developed that inhibit key processes involved in cell cycle regulation. Those approved (a) and some in clinical trials (b) shown with their respective cellular targets above. The classic anti-tumor agents are classified as: alkylating agents, and anti-metabolites, which inhibit DNA replication, topoisomerase I/II inhibitors, and mitotic spindle polymerization/depolymerization inhibitors. To date, no current cyclin D1 ablative therapies are undergoing clinical testing (Bruyère and Meijer, 2013).

1.8.1 Current FDA-Approved Targeted Therapies for RCC Renal carcinomas lack early warning signs, resulting in a significant proportion of patients initially presenting with sites of metastasis (Schöffski et al.,

2006). Patients with metastatic RCC (mRCC) have a 5-year survival rate of less than 10% (B. I. Rini et al. 2009). One problematic characteristic of RCC is its high resistance to traditional chemotherapeutics and radiotherapy, due in part to increased drug transport protein expression, rendering these treatment options largely ineffective (Cohen and McGovern 2005; Diamond et al. 2014; Siegel et al.

2014). Thus, surgical resection often represents the first line of therapy.

Moreover, immunotherapy with interleukin-2 and interferon-α (IFN-α) do not show appreciable survival advantage (Diamond et al., 2014; Escudier et al., 2007).

This points out a clear need for the continued discovery and development of targeted anti-cancer therapies for treating advanced and mRCC. Targeted therapeutics inhibiting VEGF and mTOR signaling have been developed and demonstrate significant progression free survival (PFS) in patients with advanced and mRCC. Thus, they have gained FDA approval and represent first- and second-line treatment for RCC (Coppin et al., 2011; Escudier et al., 2013; Nerich et al., 2014).

80% of human RCC cases reveal a mutation in the VHL tumor suppressor gene, most commonly arising from a deletion in chromosome 3 p (Gnarra et al.,

1996). Loss of heterozygosity at the VHL allele causes overexpression of VEGF, resulting in increased proliferation and angiogenesis. The majority of FDA

approved RCC therapies (5/7) target the VEGF signaling pathway primarily, in addition to other proteins.

Axitinib; InlytaTM (Pfizer), is a tyrosine kinase inhibitor that targets

VEGFR1-3, c-KIT, and platelet-derived growth factor receptor (PDGFR). It showed success in treating cytokine-refractory mRCC in a phase II trial (Rini et al., 2005) as well as naive mRCC in a phase II trial (Rini et al., 2013). Axitinib demonstrated higher PFS of patients with mRCC following treatment with a different, unsuccessful FDA approved systemic therapeutic (Motzer et al., 2013).

Sunitinib; SutentTM (Pfizer), is a receptor tyrosine kinase (RTK) inhibitor that targets VEGFRs, PDGFRs, KIT, CSF-1R,and flt3 (Gan et al., 2009). In a phase

III trial, sunitinib caused a higher response rate as well as longer PFS in patients with mRCC, compared to IFN-α (Motzer et al., 2007). Pazopanib; VotrientTM

(GlaxoSmithKline), is an RTK inhibitor that targets VEGFR, PDGFR, c-KIT, and fibroblast growth factor receptor (FGFR) (Pick and Nystrom, 2012; Zivi et al.,

2012). A phase III trial concluded that pazopanib demonstrated significant improvement in PFS and tumor response compared with placebo in treatment- naive and cytokine-pretreated patients with advanced and/or mRCC (Sternberg et al., 2010). Bevacizumab; AvastinTM (Genentech/Roche) is a recombinant humanized monoclonal antibody that inhibits vascular endothelial growth factor A

(VEGF-A) (Los et al. 2007). Phase II trials demonstrated a significant increase in

PFS in patients with mRCC, compared to placebo (Yang, 2004; Yang et al.,

2003).

The other group of FDA approved therapies for treating advanced and mRCC inhibit mammalian target of rapamycin (mTOR) kinase, resulting in disruption of the synthesis of proteins that regulate cell cycle, proliferation, growth and survival of tumor cells. Everolimus; AfinitorTM (Novartis) and temsirolimus; ToriselTM (Wyeth) are structurally related to sirolimus; RapamycinTM

(Wyeth), a macrolide used clinically for its immunosuppressant properties in preventing organ transplant rejection (McAlister et al., 2002). Sirolimus also shows promise in treating TSC in humans, as benign tumors in the brain, heart, skin, and kidney of TSC patients went into remission following sirolimus treatment (Dabora et al., 2011; Herry et al., 2007; Krischock et al., 2010;

Malinowska et al., 2013). Thus, everolimus and temsirolimus were developed for the treatment of RCC. Two phase III trials found that everolimus more than doubled PFS in patients with mRCC, compared to placebo (Motzer et al., 2010,

2008). A phase III trial determined that temsirolimus afforded longer overall survival as well as PFS in patients with advanced and mRCC, compared to IFN-α

(Hudes et al., 2007). The mTOR inhibitors everolimus and temsirolimus also target VEGF signaling, albeit indirectly. mTOR is important in regulating

HIFα levels, which are significantly raised in RCC due to VHL loss. In RCC, activated mTOR further increases accumulation of HIFα by increasing its synthesis which results in higher VEGF levels (Thomas et al., 2006). Thus, mTOR inhibitors are effective in decreasing cellular levels of HIFα, VEGF production and, ultimately, tumor proliferation and angiogenesis.

1.8.2 Sorafenib In addition to the above mentioned therapeutics, sorafenib; NexavarTM

(Bayer/Onyx);(4-[4-[[4-chloro-3(trifluoromethyl)phenyl]carbamoylamino]phenoxy]-

N-methyl-pyridine-2-carboxamide), is also FDA approved for the treatment of advanced and mRCC. Similar to other VEGF inhibitors, sorafenib targets

VEGFR, and PDGFR as well as C-Raf (Raf-1) and B-Raf (both wt and constitutively active B-Raf mutant V600E) (Keating and Santoro, 2009).

Sorafenib is orally active, and was the first compound in its class to reach clinical trials (Heim et al., 2005; Kohno and Pouyssegur, 2003). Sorafenib gained FDA approval in 2005 for the treatment of advanced RCC. A phase III trial found that sorafenib doubled PFS in patients with advanced RCC in whom previous therapy had failed, compared to placebo (Escudier et al., 2007).

A clinical pharmacokinetic study revealed that sorafenib is metabolized by two pathways: phase I oxidation mediated by cytochrome P450 CYP 3A4, and phase II glucuronidation mediated by the UGT1A9 pathway (Lathia et al., 2006).

The main CYP3A4 generated metabolite is sorafenib N-oxide (less than 30%).

Other minor metabolites are formed via N-methylhydroxylation and N- demethylation. The mean half-life for sorafenib ranges from approximately 25-48 hours. Following oral administration in healthy volunteers, approximately 19% of the dose was excreted in urine (mostly glucuronide conjugates of parent compound) and 77% was excreted in the feces (50 % as unchanged drug).

Neither the N-oxide metabolite nor the glucuronide metabolite of sorafenib were detected in the feces. 53

Sorafenib is taken in 200 mg tablets, twice daily, which is near the maximum tolerated dose (Awada et al., 2005). The most common side effects

(>10% of patients) are hemorrhage, hypertension, rash, fatigue, pain nausea and vomiting (Awada et al., 2005). Less common but severe side effects include leucopenia, neutropenia, thrombocytopenia, hypocalcemia, hypokalemia, congestive heart failure, myocardial infarction, QT interval prolongation, pancreatitis, gastritis and renal failure. In addition to causing negative physical effects, sorafenib can severely impact psychological well-being, evidenced by the interruption of treatment seen in some patients (Blanchet et al., 2010; Escudier et al., 2007).

1.8.3 Current RCC Adjuvant Treatment Outlook With respect to toxicity profile, sorafenib is not alone amongst the FDA approved RCC treatments. All of the above discussed treatments carry commonly observed moderate to severe side effects. Combinations of these drugs are undergoing extensive research in hopes that combination therapy will lower the effective dose of each respective drug, thus lowering the chance of adverse reactions. Currently, it is difficult to find an ongoing clinical study using only one of the above listed drugs given in monotherapy. In addition to lowering toxicity, combination therapy has the potential to raise efficacy and overcome resistance that is often seen in tumors when treated with a single agent.

Unfortunately, to date, no effective adjuvant treatment for RCC has been described (including radio-, immuno- and chemotherapies), though numerous trials involving targeted agents have not yet reported results (Janowitz et al., 54

2013). With a 5-year relapse rate for intermediate- and high-risk early-stage RCC of 30-40% (Escudier et al., 2012), research in the area of combination and adjuvant therapies is crucial. Advances in our understanding of the molecular pathogenesis of RCC will identify additional potential targets for adjuvant treatment, which should expand and diversify adjuvant treatment options.

1.9 Pentoxifylline

Pentoxifylline (PTX); TrentalTM (Sanofi); (3,7-Dihydro-3,7-dimethyl-1-(5- oxohexyl)-1H-purine-2,6-dione), a methylxanthine derivative, first gained FDA approval in 1984 (Figure 1.8). PTX is used in the treatment of intermittent claudication, vascular dementia, and other diseases involving defective locoregional blood flow (Ward and Clissold, 1987). Other indications for which the literature supports its use include: sickle cell disease, multi-infarct dementia, peripheral neuropathy, endometriosis, alcoholic and non-alcoholic steatohepatitis, and renal toxicity prevention in patients with alcoholic hepatitis and advanced chronic kidney disease (Laczy et al., 2009; Li et al., 2011; Parker et al., 2013; Sherer and Glover, 2000).

PTX has been shown to increase blood flow by decreasing fibroblast proliferation, inhibiting platelet aggregation, decreasing plasma fibrinogen concentrations, and increasing erythrocyte deformability (Aviado and Dettelbach,

1984). Its primary mechanism of action is believed to be non-specific inhibition of phosphodiesterases (PDEs), leading to accumulation of cyclic adenosine

monophosphate (cAMP) which, in erythrocytes, increases deformability, and in platelets, inhibits aggregation (Okunieff et al., 2004; Ward and Clissold, 1987).

The suggested dose of PTX is 400 mg, taken orally, three to four times a day (or 1,200 mg to 1,600 mg per day). PTX has a bioavailability of 10-30% and is extensively metabolized in the liver, as well as in erythrocytes. Less than 1% of the therapeutic dose remains unchanged in the urine. PTX has a half-life of 24-

48 min (parent drug) and 60-96 min (metabolites). Of the seven identified metabolites (M1-7) of PTX in human urine, M1 (lisofylline; Fig 1.8) and M5 posses significant hemorheologic effects while M2, M3, M4, M6 and M7 do not

(Ambrus et al. 1995). Pentoxifylline is very well tolerated. Dose-limiting side- effects (most commonly nausea) occur at approximately 2,400 mg per day. Less than 10% of patients experience adverse effects such as dizziness, nausea, and indigestion. Less that 1% experience more serious adverse effects such as angina and palpitations. PTX has minor to insignificant drug interactions with other hemorheologic therapeutics (e.g. warfarin).

PTXs utility as a palliative therapy in cancer patient care has been explored. PTX has shown promise in relieving some of the toxicities associated with various radio- and chemotherapies including cachexia, retinopathy, myelopathy, oral and intestinal mucositis, pulmonary fibrosis, and procitis/enteritis/mastitis (Bese et al., 2007; Hille et al., 2005; Kim and Park,

2006; Mantovani and Madeddu, 2010; Melo et al., 2008; Steeves and Robins,

1998; Tazi and Errihani, 2010; Theis et al., 2010; Worthington et al., 2010). The

palliative dose of PTX in cancer treatment is similar to the recommended dose for intermittent claudication (1,200 to 1,600 mg/day) and is as equally well tolerated. It is believed that PTXs hemorheologic effects play a significant role in its palliative utility, in addition to attenuating TNFα, though specific mechanisms are mostly unknown.

Such hemorheologic effects also implicate the use of PTX as a sensitizer for other anti-cancer treatments. PTX was found to decrease elevated tumor interstitial fluid pressure (TIFP) as well as increase pO2 in murine fibrosarcoma tumors, indicative of increased tumor perfusion (Lee et al. 2000). These findings demonstrate a significant ability of PTX in increasing oxygenation, systemic drug delivery, and ameliorating hypoxia in solid tumors. Of note, PTX had no observed effect on mean arterial blood pressure.

In addition to increasing tumor perfusion, PTX was discovered to increase intracellular concentrations of doxorubicin (DOX) and circumvent resistance in multidrug resistant (MDR) P388/DOX leukemia cells (Viladkar and Chitnis, 1994) as well as vincristine (VCR) in MDR L1210/VCR mouse leukemic cells (Breier et al., 1994). Multidrug resistance of both these cell lines was accompanied by overexpression of mdr1 gene product: P-glycoprotein (Pgp), a transporter responsible for the development of MDR phenotype in several neoplastic cells

(Krishna and Mayer, 2000; Kvackajová-Kisucká et al., 2001). PTX treatment caused significant Pgp downregulation in these cells via a decrease in the transcription of mdr1 (Drobná et al., 2002). It was later determined that PTX

induces the sensitization of MDR cells to VCR via downregulation of Pgp, stimulation of apoptosis and reduction of MMPs released from drug-resistant

L1210/VCR cells (Barancik et al., 2012). PTXs ability to increase tumor perfusion and reverse MDR phenotype via downregulation of pgp further nominate its use as an adjuvant therapy in cancer treatment.

Indeed, PTX significantly potentiates numerous cancer cell and tumor types to radiation therapy (Akudugu et al., 2013; Amano et al., 2005; Collingridge and Rockwell, 2000; Kinuya et al., 2001; Lee et al., n.d.; Theron et al., 2000). A phase III randomized multicenter trial investigating the effect of PTX on radiation response of patients with non-small cell lung cancer (NSCLC) concluded that

PTX was an effective radiation response modifier and provided significant benefit in the treatment of NSCLC (Kwon et al., 2000). In addition, PTX was shown to be a sensitizer for radioimmunotherapy in colon cancer xenografts (Kinuya et al.,

2001), adriamycin-induced apoptosis of leukemic cells in vitro and in vivo

(Lerma-Díaz et al., 2006), adriamycin-induced apoptosis in cervix cancer cells

(Bravo-Cuellar et al., 2010), fludarabine cytotoxicity on non-Hodgkin's lymphoma cells (Alas et al., n.d.), and cisplatin-induced apoptosis in human cervical tumor cells (Hernandez-Flores et al., 2011).

Finally, recent studies have demonstrated PTX to yield possible anticancer roles itself. PTX had antiproliferative and antimigratory effects in B16F10 melanoma cells (Dua and Gude, 2008, 2006) and xenografts (Kamran and Gude,

2013). In addition, PTX has been shown to induce apoptosis in HuT-78 human T-

cell lymphoma cells (Rishi et al., 2009) and U937 human leukemia cells (Bravo-

Cuellar et al., 2013). Further, PTX impeded migration and metastasis in MDA-

MB-231 breast cancer cells (Goel and Gude, 2013, 2011) and xenografts (Goel and Gude, 2014a), as well as A375 melanoma cells and xenografts (Kamran and

Gude, 2012). Of note, the concentration of PTX used in a majority of these studies were reported to be non-toxic and within the dose range used for the studies presented herein.

Figure 1.8 Structural comparison between cAMP, PTX and PTX metabolite 1; lisofylline The methylxanthines pentoxifylline, and its active metabolite lisofylline, compete with cAMP to non-selectively inhibit phosphodiesterases, which convert cAMP to AMP. Such inhibition raises intracellular cAMP concentrations which is believed to be responsible for the majority of pentoxifylline's and lisofylline's hemorheologic effects.

1.10 Dissertation Aims

Previous findings establish PTX to have significant palliative and sensitizing effects in clinical cancer treatment. Recent, but limited, studies also reveal PTX to illicit anticancer effects, including the inhibition of proliferation, migration and metastasis as well as induction of apoptosis in several cancer cell and xenograft models. The mechanisms behind these findings are not fully understood. Along with its robust safety profile, these anticancer effects further support PTXs utility as an adjuvant therapy in cancer treatment and warrant further understanding.

PTXs ability to decrease cyclin D1, p-Rb and arrest cancer cells in the G1 phase is not thoroughly appreciated. Therefore, in vitro studies were performed to determine the extent to which PTX could decrease cyclin D1, p-Rb and arrest

QTRRE, ACHN, 786-O and Caki-2 RCC cell models (Chapter 2; Aim 1).

Microarray results performed on PTX-treated QTRRE displayed significant G1 phase cell cycle arrest via the down-regulation of a multitude of genes involved in cell cycle progression and DNA replication. Western blot and cell cycle analysis performed on QTRRE, ACHN, Caki-2 and 786-O revealed PTXs novel ability to decrease cyclin D1 protein levels, as well as to induce G1 phase cell cycle arrest in the different RCC models, albeit to different extents.

We subsequently explored possible mechanism(s) by which PTX decreased cyclin D1 expression (Chapter 3; Aim 2). In vitro studies, mostly within the ACHN RCC cell model (due to its high sensitivity to PTX), were carried 61

out to determine how PTX caused a decrease in cyclin D1 protein levels. Results from these studies suggest that neither CCND1 transcription, nor cyclin D1 protein stability, seem to be affected by PTX treatment. Instead, results identified novel drug effects exerted by PTX, including modulation of p-4EBP1 levels at

Ser65, Thr70 and Thr 37/46, that occur with decrease in cyclin D1 and G1 phase arrest.

In vitro and in vivo ACHN xenograft studies using PTX and sorafenib were conducted to determine PTXs possible role as an adjuvant therapy via its ability to cause G1 phase arrest (Chapter 4; Aim 3). The combination of PTX + sorafenib treatment yielded significantly higher efficacy than either drug treatment alone in in vitro proliferation assays. In addition, pre-treatment with

PTX allowed for a lower effective concentration of sorafenib in these assays.

Combination PTX + sorafenib treatment yielded significantly higher tumor growth inhibition in the in vivo xenograft studies. In addition, combination drug treatment significantly decreased p-4EBP1 levels, concomitant with cyclin D1 and p-Rb levels, confirming the observed mechanism identified in the in vitro studies.

In summary, the work presented in this dissertation characterizes PTXs novel ability to decrease p-4EBP1, cyclin D1 and p-Rb levels in cancer cells and xenograft tissue, ultimately leading to G1 phase arrest. Furthermore, PTX increases the efficacy of sorafenib, as determined by cell growth inhibition and xenograft tumor growth inhibition. These data describe a desirable adjuvant

therapeutic potential for PTX, and further support its use as an adjuvant treatment in RCC, and possibly other cancers.

CHAPTER 2: PENTOXIFYLLINE DECREASES CYCLIN D1, pRB LEVELS AND ARRESTS RCC CELL MODELS IN THE G1 PHASE

2.1 Introduction

Self-sufficiency in growth signals, a hallmark of cancer, results in an enhanced cell proliferation rate and is one of the most studied aspects of cancer research (Hanahan and Weinberg, 2000). Increased cell proliferation is achieved through a multitude of mechanisms but typically involves the dysregulation of the cell cycle. Cyclin D1, a well established human oncogene, has a high frequency of amplification and overexpression in a wide variety of cancers, including breast, lung, melanoma, bladder, esophageal, colorectal, and renal, among others

(Beroukhim et al., 2010; Santarius et al., 2010). Overexpression of cyclin D1 leads to increased activity of CDK4/6, which phosphorylates and inhibits the tumor suppressor, Rb at residue Ser780 (Zheng and Lee, 2001). Rb Ser780 phosphorylation by cyclin D1-CDK4/6 causes activation of the E2F transcription factor, allowing for the transcription of genes that promote G1/S phase transition

(e.g. cyclin E) (Harbour and Dean, 2000), ultimately leading to an enhanced proliferation rate. Because of their significant role in tumorigenesis, cyclin D1 and

CDK4/6 represent logical therapeutic targets for treating cancer.

CDK-specific inhibitors are newer and less developed, compared to the multi-kinase (VEGFR, PDGFR, c-KIT, flt3, etc) inhibitors used in RCC cancer treatment. These drugs seek to inhibit cell cycle progression and are currently being investigated in both pre-clinical and clinical settings (Abate et al., 2013;

Diaz-Moralli et al., 2013; Esposito et al., 2013; Gallorini et al., 2012). Initially, pan-CDK inhibitors failed to meet expectations. However, a second generation of highly specific cyclin D1/CDK4/6 inhibitors are proving to be more efficacious in slowing the growth of tumors (Casimiro et al., 2014). Akin to CDK4/6 inhibitors, compounds that decrease cyclin D1 levels would effectively decrease CDK4/6 activity, thus slowing or arresting cell cycle progression. To date, no known cyclin

D1 ablative therapies are undergoing clinical testing.

Pentoxifylline (PTX), a non-specific phosphodiesterase inhibitor, is used as a palliative therapy in cancer patient care for its ability to relieve some of the toxicities associated with various radio- and chemotherapies (Mantovani and

Madeddu, 2010; Tazi and Errihani, 2010; Theis et al., 2010; Worthington et al.,

2010). In addition, PTX can sensitize other anti-cancer treatments, including radio- and chemotherapy in vitro and in vivo (Akudugu et al., 2013; Amano et al.,

2005; Bravo-Cuellar et al., 2010; Hernandez-Flores et al., 2011) and in patients receiving radiation treatment for non-small cell lung cancer (Kwon et al., 2000).

Recently, PTX showed antiproliferative effects in breast cancer cells (Goel and Gude, 2011), melanoma cells (Dua and Gude, 2006; Kamran and Gude,

2012; Sztiller-Sikorska et al., 2014), T-cell lymphoma cells (Rishi et al., 2009), and leukemia cells (Bravo-Cuellar et al., 2013). Mechanisms behind PTXs antiproliferative effects in these cancer models remain largely unknown, as does its ability to exert such effects in cellular models of RCC.

In the present studies, we characterized PTXs ability to decrease cyclin

D1 and induce G1 phase arrest in various RCC cell models. We began research in the QTRRE RCC cell model because of our previous and expansive work done with these cells. We then initiated research in three human RCC cell models, ACHN, Caki-2, and 786-O, that display striking similarities to the QTRRE cell model. Our results indicate that PTX significantly decreased cyclin D1 and induced G1 phase arrest in the QTRRE and ACHN RCC cell lines. This phenomenon occurred to a lesser extent in Caki-2 and 786-O, as well as the non-tumorigenic, immortalized human epithelial kidney cell model, HK-2. We further characterized PTXs effect on cell viability, as well as its ability to decrease cyclin D1, pRb Ser780, and induce G1 phase cell cycle arrest in ACHN in a time- and dose-dependent manner.

2.2 Materials and Methods

2.2.1 Cell Culture The rodent QTRRE cell line was established from primary renal epithelial cells (Yoon et al., 2001). QTRRE cells were grown in DMEM/F12 (1:1) (Corning,

Manassas, Virginia) supplemented with 10% (v/v) FBS. Human immortalized, non-tumorigenic cells, HK-2 (CRL-2190; ATCC, Manassas, Virginia), were grown in Keratinocyte-SFM media supplemented with Epidermal Growth Factor 1-53 and Bovine Pituitary Extract (Corning). Human RCC cells, Caki-2 (HTB-47;

ATCC) and 786-O (CRL-1932; ATCC), were grown in RPMI 1640 (Corning) supplemented with 10% (v/v) FBS. Human RCC cells, ACHN (CRL-1611;

ATCC), were grown in EMEM (Corning) supplemented with 10% (v/v) FBS. All cells were grown at 37ºC in a humidified atmosphere of 5% CO2. All drug treatments lasting less than 12 hrs occurred in media supplemented with 2%

(v/v) FBS.

2.2.2 Western Blot Analysis Whole cell lysates were generated using Cell Lysis Buffer 10X (Cell

Signaling Technologies, Danvers, Massachusetts) containing 1 mM Pefabloc SC,

Complete protease inhibitor cocktail tablet and phosphatase inhibitor cocktail tablet (Roche, South San Francisco, California). Nuclear and cytosolic fractions were generated using NE-PER Nuclear and Cytoplasmic Extraction Reagents

(Pierce Biotechnology, Rockford, Illinois) following manufacturer's protocol.

Protein concentration was determined using the DC Protein Assay (Bio-Rad,

Hercules, California). Protein (40µg) was subjected to 10, 12 or 14% SDS-PAGE at 140V, followed by electrophoretic transfer to PVDF membranes. Primary antibodies used were cyclin D1 (2926), pRB Ser780 (9307), SP1 (5931), tuberin

(3612), p27 (2552) (Cell Signaling Technologies) and GAPDH (8245) (Abcam,

Cambridge, Massachusetts). Secondary anti-mouse and anti-rabbit immunoglobulin conjugated with horseradish peroxidase (Santa Cruz

Biotechnology, Dallas, Texas) were used at a 1:3000 dilution in 5% milk/TBS-T.

The blots were visualized with ECL Western Blotting Detection Reagent

(Thermo, Waltham, Massachusetts) using the ChemiDoc XRS+ Imager (Bio-

Rad).

2.2.3 QTRRE Expression Microarray Analysis Total cellular RNA was extracted using the RNeasy Mini kit (Qiagen,

Valencia, California) following manufacturer's protocol and concentrations and quality were determined using absorbance at A260 and A280 following analysis on an Agilent Nanochip (Agilent, Santa Clara, California). Total RNA (500 ng) was labeled, hybridized to GeneChip Rat Gene 1.0 ST Arrays (Affymetrix, Santa

Clara, California), and read using an Affymetrix scanner according to the manufacturer’s protocols at the University of Arizona Cancer Center Genomics

Shared Service. Differential gene expression was defined as a fold change ≥ ±2.

Gene cluster analysis was performed using the DAVID Bioinformatics Database

Analysis Wizard 6.7. Gene expression changes on selected genes were validated by RT-PCR.

2.2.4 RT-PCR Total cellular RNA was extracted using the RNeasy Mini kit (Qiagen,

Valencia, California) following manufacturer's protocol and concentrations and quality were determined using absorbance at A260 and A280 following analysis on an Agilent Nanochip (Agilent, Santa Clara, California). Total RNA (2 µg) was reverse transcribed to cDNA using RevertAide First Strand cDNA synthesis kit

(Thermo Scientific, Waltham, Massachusetts) following manufacturer's protocol.

RT-PCR was performed with Maxima SYBR Green/ROX qPCR Master Mix

(Thermo Scientific, Waltham, Massachusetts), using 200 ng of cDNA template for each reaction. RT-PCR was performed on the StepOnePlus™ Real-Time PCR

System (Applied Biosystems, Grand Island, New York). Primers used were as follows: cyclin D1; 5’-gcacaacgcactttctttcc-3’ (fwd) and 5’-tccagaagggcttcaatctg-3’

(rev), cyclin B1; 5'-tgtgaaagatatctatgcttacctcag-3' (fwd) and 5'- cccagtaggtattttggtctaactg-3' (rev), cyclin B2; 5'-aaaacctcaccaagttcatcg-3' (fwd) and 5'-gagggatcgtgctgatcttc-3' (rev), cyclin A2; 5'-agtgccgctgtctctttacc-3' (fwd) and 5'-catagcatggggtgattcaa-3' (rev), cyclin F; 5'-ccgaaaaggggaattttga-3' (fwd) and 5'-ctcatccgacacagacaagc-3' (rev), RPS7; 5'-ttcgagtctggcatctctca-3' (fwd) and

5'-ttatgatggcttttcgaccac-3' (rev), GAPDH; 5'-tgggaagctggtcatcaac-3' (fwd) and 5'- gcatcaccccatttgatgtt-3' (rev) (Sigma, St. Louis, Missouri). PCR thermocycler conditions were: 95ºC for 10 min, followed by 40 cycles of 95ºC for 15 s, 60ºC for

30 s and 72ºC for 30 s. Melting curve analysis occurred following RT-PCR to verify PCR product identity and specificity. Gene specific products were

normalized to ribosomal protein S7 (RPS7) and quantified using the comparative

−ΔΔCT Ct method. Fold expression was calculated using the 2 method.

2.2.5 Cell Cycle Analysis Following drug treatment, cells were washed, trypsinized, and transferred to a 50 ml centrifuge tube and spun at 1500 RPM for 10 min at 4ºC. Media was removed, and the cells were fixed by resuspending the cell pellet with 1 ml ice- cold 70% ethanol. Samples were then stored at -20 ºC overnight. Samples were then spun at 2000 RPM for 15 minutes, and ethanol was removed. Cells were resuspended in 0.5 ml cold PBS and transferred to flow cytometer tube. 25 µl

(1/20 vol) of 10 mg/mL RNAse A and 12.5 µl (1/40 vol) of 1.6 mg/mL propidium iodide were added into each tube. Tubes were incubated at 37 ºC for 30 min, and then analyzed using the FACScanto II (BD Biosciences, San Jose, California) at the University of Arizona Cancer Center Flow Cytometry Core.

2.2.6 MTS Cell Viability Assay Cells were seeded at a density of 5.0 x 105 cells per well in 96-well plates

24 hr prior to drug treatment. Cells were then treated with PTX at the indicated times and concentrations. Following drug treatment, cells were washed twice with treatment media [DMEM (-) phenol red, (-) Na pyruvate, (+) 25mM HEPES,

(+) L-glutamine); Corning] and viability was determined by MTS assay (CellTiter

96 Non-Radioactive Cell Poliferation Assay; Promega, Madison, Wisconsin) according to manufacturer's protocol. The absorbance was determined at 490 nm in a SpectraMax M2 (Molecular Devices, Sunnyvale, CA) 96-well plate reader.

2.2.7 Statistics Data are expressed as mean averages ± standard error, unless noted otherwise. Mean values were compared using a Student's T-test, two tails, with equal variances. P-values are expressed as *p<0.05, **p<0.01, and ***p<0.001.

2.3 Results

2.3.1 PTX, but not db-cAMP or theophylline, decreases cyclin D1 in QTRRE cells

We previously reported that the cAMP/Rap1B/B-Raf/MEK/ERK 1/2 pathway modulates the expression of cyclin D1 in QTRRE cells (Cohen et al.,

2011b). Previous data suggested that increased intracellular cAMP levels would increase cyclin D1 synthesis via activation of B-Raf, and subsequently increase in CCND1 transcription. In an effort to determine the effects of increased intracellular cAMP on cyclin D1 protein levels, we treated QTRRE cells for 2, 10 and 24 hrs with dibutyryl-cAMP [db-cAMP] or PDE inhibitors, PTX, or theophylline [theo] and performed Western blot analysis (Figure 2.1). At 24 hrs, both db-cAMP and theo significantly increased cyclin D1 protein levels, as expected, whereas PTX significantly decreased cyclin D1 levels (Figure 2.1A).

Further, PTX significantly decreased cyclin D1 levels at durations below 2 hrs

(Figure 2.1B), altering cyclin D1 expression significantly faster than either db- cAMP or theo.

Figure 2.1 PTX, but not db-cAMP or theophylline, decreases cyclin D1 in QTRRE cells Whole cell lysates were generated from QTRRE cells treated with dibutyryl- cAMP [db-cAMP; 1mM], PTX (1 mg/mL), or theophylline [theo; 0.6 mg/mL] for 2, 10, and 24 hrs (A) or PTX (1.0 mg/mL) for 30, 60, 90 and 120 min (B). Equal amounts of protein were separated via SDS-PAGE and immunoblotted for cyclin D1, GAPDH.

2.3.2 PTX induces gene expression changes consistent with cell cycle arrest in QTRRE

G1 phase cell cycle arrest has been reported with PTX treatment in human kidney mesangial cells in vitro and in vivo (Lin et al., 2003), and, recently, in MDA-MB-231 human breast cells in vitro (Goel and Gude, 2013). To investigate whether cell cycle arrest was occurring, concomitant with decrease in cyclin D1 (Figure 2.1), and to explore genome-wide PTX-induced expression changes, we performed a gene expression microarray study analyzing changes in 24 hr vehicle and PTX (1.0 mg/mL) treated QTRRE cells. Table 2.1 shows the three most up- and downregulated genes, by function cluster. Significantly more downregulation of genes occurred, compared to upregulation, as evidenced by higher enrichment scores (ES). The top three clusters found upregulated following PTX treatment include genes involved in the known pharmacologic effects of PTX, including those pertaining to vasoactive/vasodilative activity (ES

2.17), cAMP-dependent activity (ES 1.89), and regulation of endothelial adhesion

(ES 1.58). The top three clusters found downregulated following PTX treatment include genes involved with the cell cycle/mitosis (ES 27.98), chromosome maintenance (ES 19.65), and DNA replication (ES 16.27).

The most upregulated genes involved in vasoactive/vasodilative and platelet adhesion processes included kininogen-1-like-1 and kininogen-1 (fold change 5.17 and 4.92, respectively (Table 2.2A). In addition, chemokine ligand

12, kininogen 2, junctional adhesion molecule 2, endothelin 1, fibrinogen beta

chain, and thrombospondin 5a were also found significantly upregulated following

PTX treatment. In addition to hemorheologic-related genes, the cAMP-specific genes PDE 4B and PDE 3B were also found upregulated (Table 2.2B).

The genes most significantly downregulated with respect to cell cycle regulation included cyclins B1, B2, A2, and F with fold change -5.5, -4.99, -4.14, and -3.78, respectively, in addition to others (Table 2.3A, Figure 2.2). Cyclin D1 showed no change in expression. Further, genes involved in chromosomal integrity and DNA replication were found to be significantly downregulated.

These included topoisomerase II a (fold change -4.50), DNA primase (fold change -2.10), as well as various DNA-dependent DNA polymerase subunits

(Table 2.3B).

In order to validate the significant downregulation of cyclins B1, B2, A2 and F, we conducted real-time quantitative PCR (RT-PCR) analysis (Figure 2.3).

Consistent with microarray analysis, PTX significantly decreased the expression of cyclin B1 to 0.09 fold (95%CI 0.08 - 0.12, p<0.01), B2 to 0.11 fold (95%CI 0.08

- 0.14, p<0.01), A2 to 0.08 fold (95%CI 0.07 - 0.10, p<0.01), and F to 0.06 fold

(95% CI 0.05 - 0.07, p<0.001), compared to untreated control. Also in agreement with microarray analysis, cyclin D1 expression (1.03 fold, 95%CI 0.84 - 1.26) was not found to be significantly altered following PTX treatment.

Table 2.1 Up- and downregulated gene cluster in QTRRE following 24 hr PTX treatment Up- and downregulated genes, by functional cluster, in 24 hr PTX (1.0 mg/mL) treated QTRRE cells, as determined by expression microarray analysis. Differential gene expression was defined as a fold change ≥ ±2. Gene cluster analysis was performed using the DAVID Bioinformatics Database Analysis Wizard 6.7.

Table 2.2 Upregulated genes in QTRRE following 24 hr PTX treatment Upregulated genes in QTRRE following 24 hr PTX (1.0 mg/mL) treatment related to vessel regulation, platelet deaggregation and platelet adhesion (A) and cAMP activity (B), as determined by expression microarray analysis. Differential gene expression was defined as a fold change ≥ 2.0.

Table 2.3 Downregulated genes in QTRRE following 24 hr PTX treatment Downregulated genes in QTRRE following 24 hr PTX (1.0 mg/mL) treatment related to cell cycle regulation (A) and chromosome integrity and DNA replication (B), as determined by expression microarray analysis. Differential gene expression was defined as a fold change ≥ 2.0.

Figure 2.2 Downregulation of cell cycle related genes in QTRRE following 24 hr PTX treatment KEGG pathway denoting cell cycle related genes found downregulated (starred) in QTRRE following 24 hr PTX (1.0 mg/mL) treatment. Differential gene expression was defined as a fold change ≥ 2.0. 79

Figure 2.3 RT-PCR validation of select cell-cycle specific, downregulated genes in QTRRE following 24 hr PTX treatment RT-PCR analysis of cyclins B1, B2, A2, F and D1. Briefly, cDNA was made from 2 µg of total RNA from 24 hr vehicle and PTX (1.0 mg/mL) treated QTRRE cells. RT-PCR was performed with Maxima SYBR Green, using 200 ng of cDNA template for each reaction. Gene specific products were normalized to ribosomal protein S7 (RPS7) and quantified using the comparative Ct method. Fold expression was calculated using the 2−ΔΔCT method. Values are means and error bars show 95% CI (n=3). P-values are expressed as **p<0.01, and ***p<0.001.

2.3.3 PTX treatment for 24 hrs causes G1 phase arrest and cyclin D1 decrease in QTRRE, Caki-2 and ACHN but not in 786-O or HK-2 RCC cell models

Following the discovery that PTX caused decreased cyclin D1 protein levels as well as decreased transcription of various genes involved in cell cycle regulation in QTRRE, we sought to test its ability to induce cell cycle arrest.

QTRRE, as well as human RCC cell models ACHN and 786-O and the non- tumorigenic, immortalized human kidney epithelial cell line, HK-2 were treated with PTX (1.0 mg/mL) for 24 hrs prior to cell cycle analysis (Figure 2.4). QTRRE and ACHN displayed enhanced G1 phase arrest of 1.70 and 1.54 fold, respectively. Along with increased G1 phase cell population, QTRRE and ACHN displayed significant decreased populations of cells in the S and G2 phase, characteristic of G1 phase arrest. The human RCC cell line, 786-O, and the human kidney epithelial cell, HK-2, showed little to no change in G1 phase arrest, nor changes in S and G2 phase cell populations.

In an effort to describe differences in PTXs ability to arrest cells in the G1 phase, we analyzed cyclin D1 protein levels in the various cell models via

Western blot, following 24 hr PTX treatment (Figure 2.5). HK-2 cells do not overexpress cyclin D1 protein levels, consistent with other non-tumorigenic cell models, and no change can be seen in cyclin D1 levels, compared to QTRRE

(Figure 2.5A). Concomitant with inducing G1 phase arrest, PTX treatment

caused significant decrease in cyclin D1 levels in the human RCC cell models,

ACHN and Caki-2, but not 786-O (Figure 2.5B). Further, Western blot analysis of these cell lines demonstrate QTRRE, ACHN, and Caki-2 to be similar, with respect to low levels of tuberin, the TSC-2 product, as well as high levels of p27, the CDK4/6 inhibitor (Figure 2.5C).

Figure 2.4 Non-tumorigenic, HK-2, and RCC cell model cell cycle comparisons following PTX treatment QTRRE, human RCC cell models, ACHN and 786-O, and the non-tumorigenic, immortalized human kidney epithelial cell line, HK-2, were treated with PTX (1.0 mg/mL) for 24 hrs prior to cell cycle analysis with propidium iodide.

Figure 2.5 Non-tumorigenic, HK-2, and RCC cell model comparisons following PTX treatment (1.0 mg/mL) for 24 hrs Following treatment with PTX (1.0 mg/mL) for 24 hrs, whole cell lysates were generated from HK-2 and QTRRE (A), the human RCC models, Caki-2, 786-O and ACHN (B), and all cell lines (C). Equal amounts of protein were separated via SDS-PAGE and immunoblotted for cyclin D1, tuberin, p27, and GAPDH.

2.3.4 Cyclin D1 protein levels recover in 786-O, but not ACHN, at 24 hrs Following observation that PTX yielded no decrease in cyclin D1 levels in

786-O at 24 hrs, compared to the other human RCC models, Caki-2 and ACHN, we sought to investigate PTXs possible effects on cyclin D1 at earlier time points

(Figure 2.6). Similar to QTRRE, PTX decreased cyclin D1 in a time-dependent manner in ACHN, causing maximal decrease at 10 and 24 hrs (Figure 2.6A).

786-O, however, displayed significant decreases in cyclin D1 following PTX treatment for 4, 8 and 12 hrs (Figure 2.6B). By 24 hrs, cyclin D1 levels in 786-O appear to fully recover.

Figure 2.6 PTX effects on cyclin D1 in ACHN and 786-O Whole cell lysates were generated from ACHN (A) and 786-O (B) cells treated with PTX (1.0 mg/mL) for the durations shown. Equal amounts of protein were separated via SDS-PAGE and immunoblotted for cyclin D1, and GAPDH.

2.3.5 PTX effects on cell viability in ACHN Because ACHN displayed higher sensitivity to PTX with respect to G1 phase cell cycle arrest (Figure 2.4) and cyclin D1 decrease (Figure 2.5 and 2.6), we decided to use this cell line for further work in characterizing PTXs ability to exert its effects. In addition, ACHN was used to determine the mechanism(s) behind PTX-induced cyclin D1 decrease (Chapter 3) and for understanding

PTXs adjuvant ability alongside sorafenib, both in vitro and in vivo (Chapter 4).

ACHN cell viability was quantified following treatment with PTX at various durations of exposure and doses by measuring relative mitochondrial dehydrogenase activity via the MTS assay. Cells treated with PTX (1.0 mg/mL) for 2, 10, and 24 hrs demonstrated no significant decrease in cell viability, compared to control (untreated) (Figure 2.7A). In addition, cells treated for 24 hrs with 0.1, 0.25, 0.5, and 1.0 mg/mL PTX also demonstrated no significant difference with respect to decreased cell viability (Figure 2.7B). Because no significant decrease in cell viability was observed, we decided to use these doses and durations for subsequent studies.

The above findings are in agreement with other studies involving similar doses of PTX. Via MTT assay, it was determined that 1.0 mg/mL PTX at 24 hrs caused insignificant decrease in cell viability in MDA-MB-231 human breast cancer cells (Goel and Gude, 2014b, 2011). Furthermore, approximately 3.0 mg/mL PTX at 24 hrs decreased cell viability by 50% in the same cells. In addition, approximately 2.5 mg/mL PTX for 24 hrs significantly decreased cell

viability in A375 human melanoma cells, as measured by MTT assay (Kamran and Gude, 2012).

Figure 2.7 Effect of PTX on mitochondrial dehydrogenase enzyme activity in ACHN cells ACHN cells were treated with PTX (1.0 mg/mL) for 2, 10, and 24 hrs (A) or for 24 hrs with 0.1, 0.25, 0.5, and 1.0 mg/mL (B). Mitochondrial dehydrogenase activity was analyzed using the CellTiter 96® Aqueous Non-Radioactive Cell Proliferation (MTS) Assay. Values are means and error bars show SD (n=3).

2.3.6 PTX decreases cyclin D1 and pRB (Ser780) levels in ACHN in a time- and dose-dependent manner

ACHN cells treated with non-toxic doses of PTX in a time- and dose- dependent manner exhibited decreased cyclin D1 and pRb levels as visualized by Western blot analysis of whole cell lysates and nuclear/cytosolic protein fractions (Figure 2.8). Cyclin D1 levels were significantly attenuated at 2, 10, and

24 hrs and pRb Ser780 levels were significantly attenuated at 10 and 24 hrs when treated with 1.0 mg/mL PTX (Figure 2.8A). On a dose-dependent scale at

24 hrs, 0.5 and 1.0 mg/mL PTX significantly decreased cyclin D1, whereas only

PTX (1.0 mg/mL) significantly decreased pRb Ser780 levels (Figure 2.8B). 1.0 mg/mL PTX treatment for 4 hrs significantly decreased cyclin D1 levels in both cytosolic and nuclear fractions, whereas pRb Ser780 levels were significantly reduced in only the nuclear fraction (Figure 2.8C).

Figure 2.8 PTX decreases cyclin D1 and pRB levels in ACHN Whole cell lysates were generated from ACHN cells treated with PTX (1.0 mg/mL) for 2, 10, and 24 hrs (A.) or for 24 hrs with 0.1, 0.25, 0.5, and 1.0 mg/mL (B.). Cytosolic and nuclear fractions were generated from ACHN cells treated with PTX (1.0 mg/mL) for 4 hrs (C.). Equal amounts of protein were separated via SDS-PAGE and immunoblotted for pRb (Ser780), cyclin D1, GAPDH (cytosolic fraction control), and SP1 (nuclear fraction control). Values are means and error bars show SE (n≥3). P-values are expressed as *p<0.05, **p<0.01, and ***p<0.001.

2.3.7 PTX induces G1-phase arrest in ACHN in a time- and dose-dependent manner ACHN cells treated with PTX at durations and doses matching those found to decrease cyclin D1 and pRb Ser780 in ACHN (Figure 2.8) exhibited significantly enhanced G1 phase cell cycle arrest as demonstrated via cell cycle analysis (Figure 2.9 and Figure 2.10). At 24 hrs, 1.0 mg/mL PTX increased the percentage of ACHN cells in the G1 phase from 55.0% (SD ± 1.5%) to 81.2%

(SD ± 0.6%), representing a 1.47 fold increase (p<0.001), compared to control

(untreated) (Figure 2.9). In addition, such treatment reduced the number of cells in the S and G2 phase to 33 percent (p<0.001) and 63 percent (p<0.001), respectively, compared to control. Little to no significant change in these populations occurred at 2 and 10 hrs of PTX treatment (1.0 mg/mL).

Similarly, PTX caused a G1 phase arrest in ACHN in a dose-dependent manner (Figure 2.9). At 24 hrs, PTX doses as low as 0.1 mg/mL caused significant G1 phase population increase. At a dose of 0.25 mg/mL, PTX increased the G1 population from 55.0% (SD ± 1.5%) to 68% (SD ± 1.4%), representing a 1.24 fold increase (p<0.001), compared to control. This same dose significantly reduced the proportion of cells in the S and G2 phase to 0.74 fold (p<0.01) and 0.64 fold (p<0.01), respectively, compared to control. Cell cycle arrest appears to be more sensitive on a time-dependent scale as opposed to dose-dependence, as the highest dose used (1.0 mg/mL) only showed significant

G1 phase arrest at 24 hrs, while lower doses (0.1 and 0.5 mg/mL), also at 24 hrs, were able to replicate similar significance in inducing G1 phase arrest.

Figure 2.9 PTX induces G1-phase arrest in ACHN in a time-dependent manner ACHN were treated with PTX (1.0 mg/mL) for 2, 10 and 24 hrs. Following treatment, cells were trypsinized, fixed and incubated with propidium iodide prior to analysis via flow cytometry. Values are means and error bars show SD (n≥3). P-values are expressed as *p<0.05, and ***p<0.001, when compared to untreated control.

Figure 2.10 PTX induces G1-phase arrest in ACHN in a dose-dependent manner ACHN were treated with PTX (24 hrs) at 0.1, 0.25, 0.5 and 1.0 mg/mL concentrations. Following treatment, cells were trypsinized, fixed and incubated with propidium iodide prior to analysis via flow cytometry. Values are means and error bars show SD (n≥3). P-values are expressed as *p<0.05, **p<0.01, and ***p<0.001, when compared to untreated control.

2.4 Discussion

Activation of the Rap1/B-Raf/MEK/ERK 1/2 pathway increased cyclin D1 expression via an increase in CCND1 transcription in the QTRRE rodent RCC cell line (Cohen et al., 2011b). In an effort to implicate increased intracellular cAMP levels, and subsequent cAMP-dependent protein kinase A (PKA) activation, in the observed increased expression of cyclin D1, we treated QTRRE with dibutyryl-cAMP (db-cAMP), theophylline (theo), and PTX. db-cAMP is a stable, membrane permeable analog of cAMP that mimics the actions of endogenous cAMP (Bartsch et al., 2003). Theo, similar to PTX, is a non-specific

PDE inhibitor. Broad inhibition of PDEs results in increased intracellular cAMP levels. As expected, db-cAMP and theo resulted in significant increases of cyclin

D1, compared to untreated control (Figure 2.1A). These findings are in agreement with a study involving treatment with lysophosphatidic acid (LPA) in

P19 embryonic carcinoma cells (Kim et al., 2012). LPA-induced proliferation in these cells was associated with increased cAMP and increased expression of cyclin D1, which was attenuated in the presence of the ERK 1/2 inhibitor, UO126.

Serendipitously, PTX treatment caused a significant and more rapid decrease in cyclin D1 expression in QTRRE cells, compared to db-cAMP and theo treatment

(Figure 2.1A, B). At the time of this discovery, only one publication demonstrated similar effects of PTX in non-tumorigenic human mesangial cells (Lin et al.,

2003).

In an effort to better understand PTXs broad effects in QTRRE cells, and to investigate whether decreased cyclin D1 levels were accompanied by cell cycle perturbation, we performed a gene expression microarray study analyzing changes in 24 hr vehicle and PTX treated cells. Broad downregulation of genes occurred, with only a few genes demonstrating significant upregulation (Table

2.1, 2.2). A majority of genes found significantly upregulated were those involved in blood vessel regulation and platelet adhesion, consistent with the known hemorheologic effects of PTX on endothelial cells. PTX modulation of genome- wide transcription rates in epithelial cells is not extensively known, however, PTX was reported to decrease the expression of integrins (αV, α2, α3, α5, and β1), and reduce cellular adhesion in B16F10 melanoma cells (Ratheesh et al., 2007) and MDA-MB-231 breast cancer cells (Goel and Gude, 2014a). PTX treatment ultimately led to decreased tumor island formation in NOD-SCID mice following injection of these cells.

Genes involved in cell cycle progression and DNA replication represented a majority of the most significantly decreased following PTX treatment (Tables

2.1, 2.3, and Figure 2.2), reflective of cell cycle arrest. Most interesting was the significant downregulation of cyclins B1, B2, A2, and F, which we validated via

RT-PCR (Figure 2.3). Surprisingly, data from the microarray and RT-PCR studies showed no change in CCND1 transcription, a suggested possible mechanism behind PTX-mediated cyclin D1 decrease first observed in mesangial cells (Lin et al., 2003).

Cell cycle analysis on the RCC cell lines, as well as the non-tumorigenic human kidney epithelial cell line, HK-2, (Figure 2.4) was performed to further investigate the cell cycle arrest suggested by the microarray study, as well as the observed cyclin D1 decrease in QTRRE cells. Our results demonstrated that PTX induced G1 phase arrest in QTRRE, and ACHN, but not 786-O or HK-2. Similar to QTRRE and ACHN cells, PTX was reported to induce G1 phase arrest in breast cancer cells (Goel and Gude, 2011), leukemia cells (Bravo-Cuellar et al.,

2013), and melanoma cells (Sztiller-Sikorska et al., 2014), though mechanisms behind G1 phase induced arrest were lacking. In a time- and dose-dependent manner, PTX decreases cyclin D1 and pRb Ser780 in whole cell, as well as nuclear and cytosolic, fractions (Figure 2.8) that are concomitant with enhanced

G1 phase cell cycle arrest in ACHN (Figures 2.9 and 2.10). The decrease in cyclin D1 in the nuclear fraction is important since nuclear localization and accumulation of cyclin D1 is critical for its oncogenicity (Kim and Diehl, 2009).

These data represent the first to effectively link decreases in cyclin D1 and pRb

Ser780 in the nucleus with PTX-mediated G1 phase arrest previously observed by others.

Consistent with G1 arrest, ACHN, Caki-2 and QTRRE cells displayed significant decreases in cyclin D1 following 24 hr PTX treatment, compared to

786-O cells (Figure 2.5B) which displayed a significant decrease in cyclin D1 at

8 hr exposure to PTX, followed by rapid recovery by 24 hrs (Figure 2.6). The differences in PTX response between individual RCC cell lines, with respect to

cyclin D1 decrease and G1 phase arrest, are of interest. Tuberin, the TSC-2 gene product found null in QTRRE cells, was low in the Caki-2 and ACHN human

RCC models (Figure 2.5C). In addition, these cells displayed high levels of the

CDK4/6 inhibitor, p27. In contrast, 786-O cells, which displayed a rapid recovery in cyclin D1 levels and insignificant G1 phase arrest, yielded high levels of tuberin and low levels of p27. Presumably, PTX sensitivity is governed by a multitude of various proteins that contribute to the regulation of cyclin D1 synthesis, intracellular localization, and stability. These results are the first to suggest that tuberin and/or p27 status may affect a particular cell-type response to PTX treatment. ACHN, but not Caki-2 or 786-O cells, are null for VHL, a situation commonly observed in human RCC (Walker, n.d.; Walker et al., 1996).

This difference suggests that disparate response to PTX in the human RCC models is independent of VHL status. In addition, HK-2 appeared unaffected by

PTX dose. HK-2 do not overexpress cyclin D1, compared to the RCC cell models

(Figure 2.5A). This suggests PTX may selectively affect cell types that overexpress cyclin D1, a characteristic of tumorigenic cells.

Here, we characterized the novel ability of PTX to induce G1 phase arrest in various RCC cell models. Further, decreased cyclin D1, and subsequently pRb

Ser780, correlated with G1 phase arrest in the ACHN, the most sensitive human

RCC cell model. Further research is needed to identify possible biomarkers used to predict relative PTX response, in addition to understanding the antiproliferative effects of PTX in cancerous and non-cancerous cells. Microarray and RT-PCR

data suggest that cyclin D1 decrease is not due to decreased transcription, as previously suggested. Understanding the mechanism(s) behind PTXs ability to decrease cyclin D1 protein levels may aid in the identification of usable biomarkers, as well as provide insight into cancer-specific antiproliferative effects induced by PTX. Therefore, we will explore the possible mechanisms by which

PTX causes cyclin D1 decrease, mostly within ACHN, in Chapter 3.

CHAPTER 3: PENTOXIFYLLINE DECREASES pAKT, pGSK-3β, p4EBP1, AND CYCLIN D1, WITHOUT AFFECTING CCND1 TRANSCRIPTION NOR CYCLIN D1 PROTEIN STABILITY

3.1 Introduction

Deregulation of cyclin D1 expression can directly contribute to some of the hallmarks of cancer, including self-sufficiency in growth signals (Hanahan and

Weinberg, 2000). Overexpression of cyclin D1 is achieved via increased synthesis (transcription and translation), and protein stability. CCND1, the gene encoding cyclin D1, is a well established human oncogene and has a high frequency of amplification and overexpression in a wide variety of cancers including breast, lung, melanoma, bladder, esophageal, colorectal, and renal, among others (Beroukhim et al., 2010; Bignell et al., 2010; Curtin et al., 2005;

Santarius et al., 2010). Hyperactivation of the MAPK signaling pathway, which controls CCND1 transcription, enhances cell-cycle progression and is found in several cancers (Dhillon et al., 2007). 4EBP1 inhibits cyclin D1 translation via binding eIF4E and inhibiting eIF4E-mediated 5' cap-dependent translation (De

Benedetti and Graff, 2004; Sonenberg and Gingras, 1998).

Hyperphosphorylation of 4EBP1 (at residues Ser 65, Thr 70, and possibly others), via upstream PI3K/AKT/mTOR signaling activation, causes decreased affinity between 4EBP1 and eIF4E, allowing for increased translation of cyclin D1 to occur (Faivre et al., 2006).

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GSK-3β phosphorylates cyclin D1 at Thr 286 causing nuclear export by

CRM1 and subsequent recognition by the E3 ubiquitin ligase, SCFFBX4/αβ-crystallin, ultimately leading to proteasomal degradation in the cytosol (Figure 1.5) (Diehl et al., 1998). Disregulation of cyclin D1 nuclear export increases its oncogenic potential (Benzeno et al., 2006), suggesting that nuclear retention of cyclin

D1, resulting from altered nuclear trafficking and proteolysis, is critical for the manifestation of its oncogenicity (Kim and Diehl, 2009). Several mutations that impair phosphorylation-dependent nuclear export of cyclin D1 have been identified in endometrial cancer (Moreno-Bueno et al., 2003) as well as in esophageal carcinoma and esophageal cancer-derived cell lines (Benzeno et al.,

2006).

Pentoxifylline (PTX) has shown antiproliferative effects in melanoma cells

(Kamran & Gude, 2012; Sztiller-Sikorska et al., 2014), and T-cell lymphoma cells

(Rishi et al., 2009). Non-toxic doses of PTX caused G1 phase arrest in B16F10 melanoma cells (Dua and Gude, 2006), MDA-MB-231 breast cancer cells (Goel and Gude, 2011) and U937 leukemia cells (Bravo-Cuellar et al., 2013). We previously reported PTXs ability to arrest QTRRE, Caki-2, and ACHN RCC cell models in the G1 phase, which correlated with decreased cyclin D1 and pRb levels (Chapter 2). Mechanism(s) behind PTX-mediated cyclin D1 decrease in these, and previously reported cancer models, remain unknown. PTX decreased pAKT, and the inhibitory phosphorylation of GSK-3β at Ser 9, which correlated with decreased cyclin D1 levels, in kidney mesangial cells (Lin et al., 2003). It was hypothesized that activated GSK-3β phosphorylated cyclin D1 at Thr 286, 101

inducing its nuclear export and proteasomal degradation, leading to overall decrease. Further, the authors hypothesized decreased pAKT led to decreased

CCND1 transcription through FKHR, a member of the forkhead transcription factors (Hutchinson et al., 2001).

In this study we sought to understand the mechanism(s) behind PTX- mediated cyclin D1 decrease, with particular focus on the most sensitive human

RCC cell model previously tested, ACHN. Our results indicate that PTX decreases translation of cyclin D1 via a decrease in pAKT and p4EBP1 levels, a novel drug effect of PTX. CCND1 transcription does not appear to be affected by

PTX, as previously hypothesized. Further, GSK-3β, as well as Thr286, do not appear to be required for PTX-mediated decrease in cyclin D1, suggesting cyclin

D1 stability is also not affected by PTX, as previously hypothesized. These novel findings bring new insights into the mechanisms of PTX action on cancer cells and have considerable relevance to the rational design of novel cyclin D1 ablative therapies for use in treating cancer.

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3.2 Methods and Materials

3.2.1 Cell Culture The rodent QTRRE cell line was established from primary renal epithelial cells (Yoon et al., 2001). QTRRE cells were grown in DMEM/F12 (1:1) (Corning,

786-O (CRL-1932; ATCC), were grown in RPMI 1640 (Corning) supplemented with 10% (v/v) FBS, and ACHN (CRL-1611; ATCC), were grown in EMEM

(Corning) supplemented with 10% (v/v) FBS. All cells were grown at 37ºC in a humidified atmosphere of 5% CO2. All drug treatments lasting less than 12 hrs occurred in media supplemented with 2% (v/v) FBS.

3.2.2 RT-PCR Total cellular RNA was extracted using the RNeasy Mini kit (Qiagen,

Valencia, California) following manufacturer's protocol and concentrations and quality were determined using absorbance at A260 and A280 following analysis on an Agilent Nanochip (Agilent, Santa Clara, California). Total RNA (2 µg) was reverse transcribed to cDNA using RevertAide First Strand cDNA synthesis kit

(Thermo Scientific, Waltham, Massachusetts) following manufacturer's protocol.

RT-PCR was performed with Maxima SYBR Green/ROX qPCR Master Mix

(Thermo Scientific, Waltham, Massachusetts), using 200 ng of cDNA template for

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each reaction. RT-PCR was performed on the StepOnePlus™ Real-Time PCR

System (Applied Biosystems, Grand Island, New York). Primers used for cyclin

D1 were 5’-gaaacttgcacaggggttgt-3’ (fwd) and 5’-atttagggggtgaggtggag-3’ (rev)

(Sigma, St. Louis, Missouri). PCR thermocycler conditions were: 95ºC for 10 min, followed by 40 cycles of 95ºC for 15 s, 60ºC for 30 s and 72ºC for 30 s. Melting curve analysis occurred following RT-PCR to verify PCR product identity and specificity. Gene specific products were normalized to GAPDH and quantified using the comparative Ct method.

3.2.3 Western Blot Analysis Whole cell lysates were generated using Cell Lysis Buffer 10X (Cell

Signaling Technologies, Danvers, Massachusetts) containing 1 mM Pefabloc SC,

(Pierce Biotechnology, Rockford, Illinois) following manufacturer's protocol.

Protein concentration was determined using the DC Protein Assay (Bio-Rad,

Hercules, California). Protein (40µg) was subjected to 10, 12 or 14% SDS-PAGE at 140V, followed by electrophoretic transfer to PVDF membranes. Primary antibodies used were: p-cyclin D1 Thr 286 (2921), cyclin D1 (2926), p-GSK-3β

Ser9 (9336), GSK-3β (9315), pAKT Thr308 (9275), pAKT Ser473 (9271), AKT

(9272), HA (3724), p-4EBP1 Ser65 (9451), Thr70 (5078) and Thr37/46 (2855) and 4EBP1 (9452) (Cell Signaling Technologies) and GAPDH (8245) (Abcam,

Cambridge, Massachusetts). Secondary anti-mouse and anti-rabbit 104

immunoglobulin conjugated with horseradish peroxidase (Santa Cruz

Biotechnology, Dallas, Texas) were used at a 1:3000 dilution in 5% milk/TBS-T.

The blots were visualized with ECL Western Blotting Detection Reagent

(Thermo, Waltham, Massachusetts) using the ChemiDoc XRS+ Imager (Bio-

Rad)..

3.2.4 Plasmids and Site-Directed Mutagenesis The Rc/CMV wt HA-cyclin D1 plasmid (8948) (Baker et al., 2005) was purchased from Addgene (Cambridge, Massachusetts). The QuickChange

Lightning Site-Directed Mutagenesis Kit (Agilent, Santa Clara, California) was used to generate the T286A HA-cyclin D1 plasmid, following manufacturer's protocol, using the wt HA-cyclin D1 as template and primers: 5'- ggacctggcttgcgcacccaccgacgtgcgggacgtgg-3' (fwd) and 5'- ccacgtcccgcacgtcggtgggtgcgcaagccaggtcc-3' (rev) (Sigma, St. Louis, Missouri)

(Thr 286 to Ala mutating codon shown in bold). Sequencing of the wt-HA and

T286A HA-cyclin D1 plasmids to validate Thr/Ala status was performed at the

University of Arizona Genetics Core using a primer for the T7-promoter. Plasmids were transformed into XL10 Gold ultracompetent cells (Agilent) following manufacturer's protocol, and purified using the Plasmid Midi Kit (Qiagen,

Valencia, California). Plasmid concentrations and quality were determined using absorbance at A260 and A280 following analysis on an Agilent Nanochip

(Agilent, Santa Clara, California).

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3.2.5 Plasmid and siRNA Transfection ACHN and HK-2 cells were seeded at 30% and 40% confluency, respectively, 24 hr prior to plasmid transfection. Wt HA-cyclin D1 and T286A HA- cyclin D1 plasmids were transfected into cells using Lipofectamine LTX with

PLUS Reagent (Life Technologies, Grand Island, New York) following manufacturer's protocol. Opti-MEM I reduced-serum media (Life Technologies) and serum-free Keratinocyte-SFM media (Corning, Manassas, Virginia) were used in transfection of ACHN and HK-2 cells, respectively. All media used in transfections were supplied by Life Technologies (Grand Island, New York).

ACHN and HK-2 cells were seeded at 30% and 40% confluency, respectively, 24 hr prior to transfection. SMARTpool ON-TARGETplus GSK-3β and siGENOME non-targeting control.siRNA (Dharmacon RNAi Technologies, Lafayette,

Colorado) were transfected into cells using DharmaFECT 1 Transfection

Reagent (Dharmacon), according to manufacturer's protocol. All siRNA and siRNA transfection reagent were supplied by Thermo Scientific (Rockford, IL).

3.2.6 Immunoprecipitation ACHN total cell lysate (1 mg) tumbled with 10 µg cyclin D1 (2926) antibody (Cell Signaling Technologies, Danvers, Massachusetts) for 1 hr at 4 ºC.

25 µL protein G sepharose bead slurry (Amersham, Pittsburgh, PA) was added and incubated with rotation overnight at 4 ºC. The beads were washed thrice with Cell Lysis Buffer (Cell Signaling Technologies) containing 1 mM Pefabloc

SC, Complete protease inhibitor cocktail tablet and phosphatase inhibitor cocktail tablet (Roche, South San Francisco, California). 4x XT sample loading buffer 106

(Bio-Rad) with 5% β-mercaptoethanol (v/v) was added to the beads and boiled for 5 min to elute the proteins in preparation for Western blot analysis.

3.2.7 Statistics Data are expressed as mean averages ± standard error, unless otherwise stated. Mean values were compared using a Student's T-test, two tails, with equal variances. P-values are expressed as *p<0.05, **p<0.01, and ***p<0.001.

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3.3 Results

3.3.1 PTX has no effect on CCND1 transcription rates in QTRRE, ACHN or 786-O

A decrease in CCND1 transcription was previously suggested to be a factor in the first observation of decreases in cyclin D1 following PTX treatment in kidney mesangial cells stimulated with PDGF (Lin et al., 2003). To explain the mechanism behind PTX-mediated cyclin D1 decrease seen previously in our

RCC cell models (Figures 2.1, 2.5, 2.6 and 2.8), we first analyzed cyclin D1 mRNA copy number via RT-PCR using similar PTX doses and durations. With respect to time, PTX treatment (1.0 mg/mL) had no affect on relative Ct values in

QTRRE, ACHN and 786-O, compared to untreated control (Figure 3.1 A, C and

E, respectively). Further, at 24 hrs of PTX treatment (ranging from 0.1 to 1.0 mg/mL), no significant change in Ct was observed in QTRRE or ACHN, compared to untreated control (Figure 3.1 B and D, respectively).

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Figure 3.1 PTX is without effect on CCND1 transcription rates mRNA was isolated from QTRRE (A and B), ACHN (C and D), and 786-O (E) cells treated with PTX (1.0 mg/mL) for the durations shown or for 24 hrs with the concentrations shown. Equal amounts of cDNA were analyzed via RT-PCR. Gene specific products were normalized to GAPDH and quantified using the comparative Ct method. Values are means and error bars show SD (n≥3).

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3.3.2 PTX decreases pAKT (Thr308 and Ser473), p4EBP1 (Ser 65, Thr 70, and Thr 37/46) and cyclin D1 levels in a time- and dose- dependent manner in ACHN

Ruling out changes in the transcription of cyclin D1, we next examined modulation of the PI3K/AKT/mTOR/4EBP1 pathway, responsible for cyclin D1 translation. PTX (1.0 mg/mL) decreased pAKT Thr 308 and Ser 473 levels at 2 and 10 hrs, followed by recovery of these levels (Figure 3.2). AKT phosphorylates GSK-3β at Ser 9, thus we also investigated its phosphorylation status following PTX treatment. Parallel to decreased pAKT, pGSK-3β levels were significantly attenuated following 2 hrs PTX treatment, followed by recovery at 10 and 24 hrs (Figure 3.2). Also downstream of AKT is mTOR-mediated sequential phosphorylation of the translation inhibitor, 4EBP1 (Ser 65, Thr 70, and Thr 37/46). Parallel with pAKT loss, PTX caused decreased levels of p4EBP1 at Ser 65, Thr 70, and Thr 37/46 at 2, 10, and 24 hrs treatment, compared to untreated control (p<0.05) (Figure 3.3A). PTX (1.0 mg/mL) treatment at 24 hrs (compared to 0.1, 0.25, and 0.5 mg/mL) significantly decreased p4EBP1 levels, compared to untreated control (p<0.05) (Figure

3.3B). Total 4EBP1 displays a decrease in molecular weight following these drug treatments, consistent with loss of post-translational modifications.

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Figure 3.2 PTX decreases pAKT and pGSK-3β levels in ACHN cells

Whole cell lysates were generated from ACHN cells treated with PTX (1.0 mg/mL) for 2, 10, and 24 hrs. Equal amounts of protein were separated via SDS- PAGE and immunoblotted for pAKT (Thr308 and Ser473), pGSK-3β (Ser9), GSK-3β, and GAPDH. Values are means and error bars show SE. P-values are expressed as **p<0.01, as compared to untreated control.

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Figure 3.3 PTX decreases p-4EBP1 levels, concomitant with cyclin D1 decrease in ACHN cells Whole cell lysates were generated from ACHN cells treated with PTX (1.0 mg/mL) for 2, 10, and 24 hrs (A) or for 24 hrs with 0.1, 0.25, 0.5, and 1.0 mg/mL (B). Equal amounts of protein were separated via SDS-PAGE and immunoblotted for cyclin D1, p4EBP1 Ser65, Thr70, Thr37/46, total 4EBP1, and GAPDH. Values are means and error bars show SE (n≥3). P-values are expressed as *p<0.05, compared to untreated control.

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3.3.3 PTX-mediated cyclin D1 decrease is independent of GSK-3β activity

PTX decreased pAKT, and the inhibitory phosphorylation of GSK-3β at

Ser 9, which correlated with decreased cyclin D1 levels in kidney mesangial cells

(Lin et al., 2003). It was hypothesized that activated GSK-3β phosphorylated cyclin D1 at Thr 286, inducing its nuclear export and proteasomal degradation, leading to an overall decrease. Consistent with this finding, we observed decreased pAKT and inhibitory pGSK-3β levels following PTX dose (Figure 3.2).

To investigate any pharmacological and genetic manipulation on PTX- mediated cyclin D1 decrease by GSK-3β, we used small molecule inhibitors as well as knockdown via siRNA. LiCl is a commonly used inhibitor of GSK-3β. 24 hr treatment of LiCl (10mM) induced auto-inhibitory Ser 9 phosphorylation on GSK-

3β (Figure 3.4A). Following 24 hr treatment with combination LiCl and PTX (1.0 mg/mL), cyclin D1 decrease was observed in ACHN. Further, using 20mM LiCl and another proprietary GSK-3β inhibitor, VI (10 μM; Santa Cruz Biotechnology), dual inhibition of GSK-3β failed to significantly rescue decreases in cyclin D1 at 4 and 8 hrs PTX treatment (1.0 mg/mL) in ACHN (Figure 3.4B). In addition, we used siRNA to knockdown GSK-3β levels. Following transfection with control siRNA [con] or GSK-3β siRNA, 24 hr PTX treatment (1.0 mg/mL) effectively induced cyclin D1 decrease in ACHN (Figure 3.5).

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Figure 3.4 GSK-3β inhibition via LiCl and does not affect PTX-mediated cyclin D1 decrease in ACHN cells

Whole cell lysates were generated from ACHN treated with GSK-3β inhibitor, LiCl (10 mM), PTX (1.0 mg/mL), or both for 24 hr (A), or GSK-3β inhibitors, LiCl (20 mM) and VI (10 µM; Santa Cruz Biotechnology), PTX (1 mg/mL), or both for 4 and 8 hrs (B). Equal amounts of protein were separated via SDS-PAGE and immunoblotted for, cyclin D1, and GAPDH.

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Figure 3.5 GSK-3β knockout via siRNA does not affect PTX-mediated cyclin D1 decrease in ACHN cells

Whole cell lysates were generated from ACHN, untreated or treated with PTX (1.0 mg/mL) for 24 hr, following transfection with control siRNA [con; 25 nM] or GSK-3β siRNA (25 nM). Equal amounts of protein were separated via SDS- PAGE and immunoblotted for GSK-3β, cyclin D1, and GAPDH. Values are means and error bars show SE (n=4). ns: not significant.

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3.3.4 PTX-mediated decrease in cyclin D1 is independent of DYRK 1B, and does not increase Thr 286 phosphorylation on cyclin D1

Pham=rmacological and genetic manipulation of GSK-3β failed to prevent

PTX-mediated cyclin D1 decrease (Figures 3.4 and 3.5), however, the possibility of a different kinase capable of cyclin D1 Thr286 phosphorylation following PTX treatment may exist. DYRK1B, a dual-specificity kinase, was found to phosphorylate cyclin D1 at Thr 286, driving its proteasomal turnover and inducing

G1 phase arrest in HEK-293 and PANC-1 cells, independent of GSK-3β (Ashford et al., 2014). To investigate whether DYRK1B was involved in the increased turnover of cyclin D1 due to PTX treatment, we used the novel DYRK1B inhibitor,

AZ191, using concentrations and durations similar to previous studies. Pre- treatment with AZ191 (1 or 3 µM) failed to rescue decrease in cyclin D1 caused by PTX (1.0 mg/mL) at 4 hrs (Figure 3.6).

In addition to GSK-3β and DYRK1B, other kinases promote phosphorylation on cyclin D1 at Thr 286. The MAPK protein, p38, phosphorylates cyclin D1 at Thr 286 and induce its proteasomal degradation in response to osmotic shock in Granta 519 cells (Casanovas et al., 2000). Further, p38 caused proteasomal degradation of cyclin D1 and cell cycle arrest in response to aspirin treatment in colorectal cancer cells (Thoms et al., 2007). The ataxia telangiectasia mutated (ATM) and the ATM- related (ATR) kinases were also discovered to be able to promote the phosphorylation of cyclin D1 Thr286, following activation via UV irradiation or double-stranded DNA (dsDNA) breakage

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in cells (Hitomi et al., 2008). In addition, DYRK1A, a family member of DYRK1B, was recently reported to phosphorylate Thr 286 and induce G1 phase cell cycle arrest, leading to cell cycle exit and differentiation in neuronal cells (Soppa et al.,

2014).

To test relative p-cyclin D1 Thr 286 levels, we treated ACHN between 0 and 4 hrs with MG-132 (10 µM), and combination MG-132 + PTX (1.0 mg/mL).

Because PTX leads to total cyclin D1 loss, we treated cells with the proteasome inhibitor, MG-132, to prevent cyclin D1 loss in hopes of seeing modulation of Thr

286 phosphorylation. Phosphorylation at Thr286 increased with MG-132 dose from 0 to 2 hrs (Figure 3.7A) and 0 to 4 hrs (Figure 3.7B), concomitant with an increased total cyclin D1 levels, compared to untreated control. Combination MG-

132 + PTX failed to yield significantly increased phosphorylation at Thr 286, compared to MG-132 alone, from 0 to 2 hrs (Figure 3.7A) and 0 to 4 hrs (Figure

3.7B), compared to untreated control, suggesting no increase of Thr 286 phosphorylation was occurring with PTX dose.

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Figure 3.6 Inhibition of DYRK1B via AZ191 does not affect PTX-mediated cyclin D1 decrease in ACHN cells Whole cell lysates were generated from ACHN, pre-treated with AZ191 (1 or 3 µM) for 24 hr, followed by treatment with PTX (1 mg/mL) for 4 hr. Equal amounts of protein were separated via SDS-PAGE and immunoblotted for cyclin D1, and α-tubulin.

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Figure 3.7 PTX does not increase p-cyclin D1 Thr 286 levels following 2 hr and 4 hr doses

Whole cell lysates were generated from ACHN cells, treated with MG-132 (10 µM) alone and in combination with PTX (1 mg/mL) for up to 2 hr (A) and 4 hr (B). Equal amounts of protein were separated via SDS-PAGE and immunoblotted for p-cyclin D1 (Thr 286), cyclin D1, and GAPDH. Values are means and error bars show SE (n=3).

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3.3.5 PTX-mediated cyclin D1 decrease is independent of Thr 286 phosphorylation

To further investigate whether Thr 286 was required for PTX-mediated decrease using constructs for C-terminal HA-tagged wild-type (wt) and threonine

286 to alanine mutant (T286A) forms of cyclin D1. Research using T286A, which cannot be phosphorylated by GSK-3β, or other kinases, effectively demonstrate that CRM1 nuclear export and SCFFBX4/αβ-crystallin-mediated ubiquitination are crucial in limiting cyclin D1s oncogenic potential. T286A remained nuclear throughout the cell cycle and overexpression resulted in cellular transformation in murine fibroblasts and promoted tumor growth in immune compromised mice (Alt et al., 2000). Further, transgenic mice that overexpress T286A developed mammary adenocarcinoma with a shorter latency relative to mice overexpressing the wild-type cyclin D1 (Lin et al., 2008).

We transfected ACHN with wt and T286A constructs and pulled down via immunoprecipitation against HA, following 4 hr MG-132 (10 µM) treatment to allow for possible phosphorylation at Thr 286 to accumulate (Figure 3.8A). The wt sample is positive for Thr 286 antibody recognition, compared to the T286A mutant [Mut], which shows no signal via Western blot. In addition, input HA signal is higher in the T286A sample, demonstrating increased expression of T286A compared to wt. Using the translation inhibitor, cycloheximide (CHX), we measured relative rates of degradation between wt and T286A (Figure 3.8B).

T286A is more stable, compared to wt, reflecting decreased degradation due to

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mutation at the critical Thr site. This finding is in agreement with the increased signal of HA following IP in Figure 3.8A. Finally, following transfection with

T286A mutant, we measured HA levels in response to 2, 10 and 24 hr PTX treatment (1.0 mg/mL) (Figure 3.8C). 24 hr PTX treatment caused decrease of

T286A, compared to untreated control, in ACHN cells.

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Figure 3.8 Cyclin D1-HA mutant T286A decreases with PTX dose in ACHN cells

Immunoprecipitations for HA-tag from ACHN transfected with control plasmid (empty vector, [con]), wild type-HA cyclin D1 [WT], and T286A-HA mutant [Mut], treated with MG-132 (10 µM) for 4 hrs (A). Images are derived from the same blot, lanes were removed for clarity. (B) Whole cell lysates were generated from ACHN transfected with WT and Mut, and treated with CHX from 0 to 4 hrs. (C) Whole cell lystaes were generated from ACHN transfected with Mut and treated with PTX (1.0 mg/mL) for 2, 10 and 24 hrs. Equal amounts of protein were separated via SDS-PAGE and immunoblotted for p-cyclin D1 (Thr 286), HA-tag, and GAPDH.

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3.3.6 PTX-mediated cyclin D1 decrease is independent of proteasomal degradation and does not affect cyclin D1 stability

Proteasomal degradation of cyclin D1, independent of GSK-3β activity as well as p-cyclin D1 Thr 286 status, has been reported. Antizyme (AZ) is known to be a regulator of polyamine metabolism that inhibits ornithine decarboxylase activity and polyamine transport. When treated with N-nitrosomethylbenzylamine, mice overexpressing AZ had significantly lower forestomach tumor incidence and tumor multiplicity, compared to Wt littermates (Fong et al., 2003). Further, proliferative forestomach epithelium derived from Wt mice showed overexpression of cyclin D1, compared to forestomach epithelium from mice overexpressing AZ which displayed weak staining for cyclin D1. It was discovered that AZ was capable of specific, noncovalent association with cyclin

D1 and that this interaction accelerated cyclin D1 degradation in vitro in the presence of only cyclin D1, purified 26 S proteasomes, and ATP (Newman et al.,

2004). Further, AZ overexpression caused proteasomal degradation of both wt and T286A mutant forms of cyclin D1 in rat hepatoma cells. These results suggest AZ controls the proteasomal degradation of cyclin D1, that is independent of GSK-3β, Thr286, cyclin D1s E3 ubiquitin ligase, SCFFBX4/αβ-crystallin, and ubiquitin.

To rule out whether increased proteasomal degradation of cyclin D1 was occurring, independent of GSK-3β activity and Thr 286 phospho-status, we used the 26S proteasome inhibitor, MG-132. MG-132 (10 µM) partially rescued cyclin

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D1 levels following 2 and 4 hr doses of PTX (1.0 mg/mL) in QTRRE, 786-O, and

ACHN cells (Figure 3.9 A, B, and C, respectively), however, this rescue was significantly lower when compared to MG-132 alone. Combination MG-132 and

PTX treatment led to a rescue of 65.8% (p<0.05) and 45.9% (p<0.05) at 2 and 4 hrs, respectively, compared to MG-132 alone treatment in ACHN (Figure 3.9C).

Further ruling out proteasomal degradation, we used the translation inhibitor, CHX (10 µM), to determine the relative half-life of cyclin D1 protein in

ACHN when treated with PTX (1.0 mg/mL) from 0 to 2 hrs (Figure 3.10). CHX and CHX + PTX treatment equally decreased cyclin D1 protein amounts, causing no significant change it cyclin D1 T1/2 further ruling out PTX-mediated decrease of cyclin D1 relies on proteasomal degradation, independent of GSK-3β or Thr

286 status.

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Figure 3.9 MG-132 fails to completely rescue PTX mediated cyclin D1 decrease

Whole cell lysates were generated from QTRRE (A), 786-O (B), and ACHN (C) cells treated with either the proteasome inhibitor, MG-132 (10 µM), PTX (1.0 mg/mL) or a combination of the two for 2 and 4 hrs. Equal amounts of protein were separated via SDS-PAGE and immunoblotted for cyclin D1, and GAPDH. Values are means and error bars show SE (n=3). P-values are expressed as *p<0.05

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Figure 3.10 PTX has no effect on cyclin D1 stability, as demonstrated by

T1/2

Whole cell lysates were generated from ACHN cells treated with either the translation inhibitor, CHX (10 µM), PTX (1.0 mg/mL) or a combination of the two from 0 to 2 hrs (A). Equal amounts of protein were separated via SDS-PAGE and immunoblotted for cyclin D1, and GAPDH. B Densitometry data plotted logarithmically on the y-axis for T1/2 estimation. Values are means and error bars show SE (n=3). P-values are expressed as *p<0.05.

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3.4 Discussion

Our finding that PTX had no affect on CCND1 transcription rates in

QTRRE (Figure 3.1A and B) is consistent with our previous microarray data and

RT-PCR validation showing no change in CCND1 transcription in QTRRE cells treated with PTX (1.0 mg/mL) for 24 hrs (Result 2.3.2). We also observed no changes in CCND1 transcription rates in ACHN (Figure 3.1C and D), as well as

786-O (Figure 3.1E) following PTX treatment. Taken together, this suggests that

PTX-mediated decrease of cyclin D1 protein levels in these cells is not due to decreases in CCND1 transcription, as previously hypothesized. 786-O demonstrated maximal cyclin D1 decrease at 8 hr PTX treatment (1.0 mg/mL), followed by full recovery at 24 hrs (Figure 2.6). This data also suggests that the recovery of cyclin D1 seen in 786-O is not due to an increase in CCND1 transcription.

Our results showing PTX promoted pAKT inhibition (Figure 3.2) is consistent with previously published studies. PTX inhibited the PDGF-activated phosphorylation of AKT in rat airway smooth muscle cells (Chiou et al., 2006) as well as AKT activation in fMLP-stimulated human neuronal cells (Costantini et al.,

2010). Further, PTX lowered the level of activated AKT in MDA-MB-231 human breast cancer cells (Goel and Gude, 2013). Downstream of AKT, we found PTX caused decreased p4EBP1 levels at Ser 65, Thr 70 and Thr 37/46, among other possible residues, that was time- and dose-dependent (Figure 3.3). These results represent novel PTX drug effects, and correlate decreased pAKT with

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decreased cyclin D1 protein levels, as previously observed (Lin et al., 2003).

Between pAKT and p4EBP1 is the kinase, mTOR. Presumably, PTX promotes mTOR kinase inhibition via a decrease in pAKT levels, as suggested by decreased p4EBP1 levels (Figure 3.3). Indeed, PTX has been shown to prevent proliferation and activation of mTOR in cultured spinal astrocytes (Norsted

Gregory et al., 2013). This effect essentially mimics the action of the FDA approved RCC targeted therapies, everolimus and temsirolimus, that inhibit mTOR, effectively reducing 4EBP1 phosphorylation and subsequent translation of proteins, like cyclin D1, believed to contribute to cancer progression (Faivre et al., 2006; Jastrzebski et al., 2007).

Our finding that PTX promoted inhibitory pGSK-3β Ser 9 loss (Figure 3.2) agreed with a study that found PTX decreased pAKT and pGSK-3β at Ser 9, which correlated with decreased cyclin D1 levels in kidney mesangial cells (Lin et al., 2003). The authors of this study hypothesized that activated GSK-3β phosphorylated cyclin D1 at Thr 286, inducing its nuclear export and proteasomal degradation, leading to overall cellular decrease. Small molecule inhibition as well as knockdown via siRNA rule out GSK-3β activity in being involved in the mechanism behind PTXs cyclin D1 decrease (Figures 3.4 and 3.5). DYRK1B was also found to phosphorylate cyclin D1 at Thr 286, driving its proteasomal turnover and inducing G1 phase arrest in HEK-293 and PANC-1 cells, independent of GSK-3β (Ashford et al., 2014). Small molecule inhibition of

DYRK1B demonstrate it, too, was not involved in PTX-mediated cyclin D1 decrease (Figure 3.6). Further, we observed no significant increase in 128

phosphorylation of residue Thr 286 on cyclin D1 following 2 and 4 hr PTX treatment durations (Figure 3.7). Taken together, this data is the first to suggest that GSK-3β or DYRK1B kinase activity is not required for PTX-mediated cyclin

D1 decrease.

Because we observed no overall phosphorylation increase of Thr 286 following PTX treatment, we asked whether Thr 286 was required for cyclin D1 decrease caused by PTX. The HA-T286A cyclin d1 mutant, which cannot be phosphorylated, or readily degraded, demonstrated higher stability compared to

HA-wt cyclin D1 when transfected into ACHN (Figure 3.8A and B). We found

PTX retained its ability to decrease T286A following 24 hr PTX treatment at 1.0 mg/mL (Figure 3.8C), further suggesting increased turnover of cyclin D1 is not occurring.

Finally, using the proteasome inhibitor, MG-132, we were unable to rescue cyclin D1 decrease with PTX treatment (Figure 3.9), suggesting a mechanism other than proteasomal degradation was occurring. To further rule out decreased cyclin D1 stability via other forms of degradation (lysosomal), we used the translation inhibitor CHX. CHX + PTX treatment did not cause significant decrease in cyclin D1 T1/2, compared to CHX alone (Figure 3.10), affirming an increase in cyclin D1 turnover is not occurring with PTX treatment in ACHN cells.

Cyclin D1 overexpression is a common event in cancer but does not occur solely as a consequence of gene amplification. Rather, increased levels of cyclin

D1 frequently result from its defective regulation at the post-translational level, 129

primarily in processes involved with Thr 286 phosphorylation, nuclear export, and ubiquitin-mediated proteasomal degradation (Gillett et al., 1994; Russell et al.,

1999). Indeed, several mutations that impair phosphorylation-dependent nuclear export of cyclin D1 have been identified in endometrial cancer (Ikeda et al., 2013;

Moreno-Bueno et al., 2003) as well as in esophageal carcinoma and esophageal cancer-derived cell lines (Benzeno et al., 2006). Further, high levels of cyclin D1b

(lacking the C-terminal Thr286 residue) have been reported to be associated with a poor prognosis for breast cancer patients (Abramson et al., 2010; Asiedu et al.,

2014; Taneja et al., 2010; Zhu et al., 2010) as well as non-small cell lung cancer patients (Li et al., 2008). Because PTX appears to inhibit the translation of cyclin

D1 through decreased pAKT/p4EBP1 levels (Figures 3.2 and 3.3), PTX treatment will presumably be effective in tumors containing these mutated forms of cyclin D1. Indeed, PTX treatment caused reduced protein levels of T286A in

ACHN (Figure 3.8C).

Studies are ongoing in our laboratory to identify the unknown targets regulating pAKT loss via PTX treatment. PI3K activates AKT via generation of phosphatidylinositol (3,4,5)-trisphosphate (PIP3), a phospholipid that resides on the plasma membrane. AKT is recruited to the membrane via its PH domain and is phosphorylated by phosphoinositide-dependent kinase-1 (PDK1), and other kinases, for full activation (Franke et al., 1997). The tumor suppressor, phosphatase and tensin homolog (PTEN), causes reduction in pAKT levels by converting PIP3 to PIP2 via removal of a phosphate group. PTEN is mutated or deleted in several types of cancer, namely prostate cancer, which leads to 130

increased pAKT signaling and tumor progression (Cairns et al., 1997; Whang et al., 1998). PTEN protein and gene expression have been described as reduced

(Hara et al., 2005; Shin Lee et al., 2003), absent (Brenner et al., 2002), mutated

(Alimov et al., n.d.), or deleted (Sükösd et al., 2001) in human RCC. Caffeine and pentoxifylline radiosensitized PTEN-deficient but not PTEN-proficient glioma cells, which correlated with the activation of the G1 DNA damage checkpoint and inhibition of AKT hyperphosphorylation (Sinn et al., 2010). Interestingly, ACHN contain wt PTEN (Lin et al., 2007), suggesting these effects are independent of

PTEN-status in RCC cell models. Either way, understanding the mechanisms leading to pAKT loss will help to identify PTXs ultimate target with respect to cyclin D1 decrease and subsequent G1 phase arrest in cancer cells.

These novel findings have considerable relevance to the rational design of novel chemotherapeutic and chemopreventative strategies. PTX-mediated decrease of cyclin D1 via decreased pAKT and p4EBP1 levels nominate it as an adjuvant therapy in preventing or slowing tumor cell proliferation. Therefore, we will explore the utility of PTX, in combination with the FDA approved RCC targeted therapy, sorafenib, to increase antiproliferative effects within ACHN cells and xenografts in Chapter 4.

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CHAPTER 4: PENTOXIFYLLINE INCREASES THE EFFICACY OF SORAFENIB IN ACHN CELLS AND XENOGRAFTS IN MICE

4.1 Introduction

Sorafenib; NexavarTM (Bayer/Onyx); (4-[4-[[4-chloro-3-

(trifluoromethyl)phenyl]carbamoylamino]phenoxy]-N-methyl-pyridine-2- carboxamide), is an FDA approved second-line targeted therapy for the treatment of advanced and metastatic RCC. Sorafenib inhibits VEGFR, PDGFR,

C-Raf (Raf-1), and B-Raf (both wt and constitutively active B-Raf mutant V600E)

(Keating and Santoro, 2009). Sorafenib is orally active, and was the first compound in its class to reach clinical trials (Heim et al., 2005; Kohno and

Pouyssegur, 2003). Previous data from our laboratory demonstrated that sorafenib caused decrease in cyclin D1 expression, presumably through inhibition of the Raf/MEK/ERK 1/2 signaling pathway, in the rodent QTRRE RCC cell model (Cohen et al., 2011b). Further, treatment of these cells with sorafenib produced a significant decrease in the protein expression of p4EBP1 at Ser 65,

Thr 70 and Thr 37/46, as well as cyclin D1 levels (Cohen, et al., 2011).

PTX has been demonstrated to yield antiproliferative effects in melanoma cells (Kamran & Gude, 2012; Sztiller-Sikorska et al., 2014), and T-cell lymphoma cells (Rishi et al., 2009). Non-toxic doses of PTX caused G1 phase arrest in

B16F10 melanoma cells (Dua and Gude, 2006), MDA-MB-231 breast cancer cells (Goel and Gude, 2011) and U937 leukemia cells (Bravo-Cuellar et al.,

2013). Previously, we have demonstrated PTX to also decrease cyclin D1 and 132

induce G1 phase arrest in rodent and human RCC cell models (Chapters 2 and

3). Literature further supports PTXs utility as an antiproliferative agent, as demonstrated in various xenograft models. PTX (40 and 60 mg/kg) inhibited

A375 melanoma tumor growth by 35 and 50% in xenograft models (Kamran and

Gude, 2012). In addition, PTX (40 and 60 mg/kg) were found to inhibit MDA-MB-

231 tumor growth, equally, by 36% in xenograft models (Goel and Gude, 2013).

Further, combination treatment with PTX and doxorubicin (DOX) delayed tumor growth more than either treatment alone in breast cancer xenograft models (Goel and Gude, 2014b). Mechanisms explaining PTXs antitumor effects in these xenograft models are lacking. In addition, whether PTX exerts these effects in

RCC xenografts models is unknown.

Our previous data indicate that PTX decreases translation of cyclin D1 via a decrease in pAKT and p4EBP1 levels in ACHN cells (Chapter 3). In this study we evaluated the utility of PTX to serve as an adjuvant treatment alongside sorafenib in ACHN cells as well as subcutaneous xenograft tissue. Our results reveal that PTX increases the efficacy, and lowers the effective dose, of sorafenib in ACHN cells. Combination treatment with PTX and sorafenib in

ACHN also significantly reduced cyclin D1 levels. In addition, combination treatment with PTX and sorafenib significantly inhibited tumor growth inhibition, as well as p4EBP1 and cyclin D1 protein levels in ACHN xenograft tissue, validating our previous findings in cells. These results further strengthen the possible adjuvant role that PTX may serve, alongside sorafenib and other targeted therapies, in treating RCC and other cancers. 133

4.2 Materials and Methods

4.2.1 Reagents Pentoxifylline (PTX) used in the in vivo xenograft study was purchased from Western Medical Supply (Arcadia, California) while PTX used for cell culture work was purchased from Sigma (St. Louis, Missouri). Sorafenib was purchased from Selleck Chemicals (Houston, Texas). Cremophor EL was purchased from

Sigma.

4.2.2 Cell Culture Human RCC cells, ACHN (CRL-1611; ATCC, Manassas, Virginia), were grown in EMEM (Corning, Manassas, Virginia) supplemented with 10% (v/v)

FBS. Cells were grown at 37ºC in a humidified atmosphere of 5% CO2. All drug treatments lasting less than 12 hrs occurred in media supplemented with 2%

(v/v) FBS.

4.2.3 MTS Cell Viability Assay Cells were seeded at a density of 5.0 x 105 cells per well in 96-well plates

24 hr prior to drug treatment. Cells were then treated with PTX and/or sorafenib at the indicated times and concentrations. Following drug treatment, cells were washed twice with treatment media (DMEM (-) phenol red, (-) Na pyruvate, (+)

25mM HEPES, (+) L-glutamine) (Corning, Manassas, Virgina) and viability was determined by MTS assay (CellTiter 96 Non-Radioactive Cell Poliferation Assay;

Promega, Madison, Wisconsin) according to manufacturer's protocol. The

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absorbance was determined at 490 nm in a SpectraMax M2 (Molecular Devices,

Sunnyvale, CA) 96-well plate reader.

4.2.4 Western Blot Analysis Whole cell and xenograft tissue lysates were generated using Cell Lysis

Buffer 10X (Cell Signaling Technologies, Danvers, Massachusetts) containing 1 mM Pefabloc SC, Complete protease inhibitor cocktail tablet and phosphatase inhibitor cocktail tablet (Roche, South San Francisco, California). Protein concentration was determined using the DC Protein Assay (Bio-Rad, Hercules,

California). 40µg protein was subjected to 10, 12 or 14% SDS-PAGE at 140V, followed by electrophoretic transfer to PVDF membranes. Primary antibodies used were cyclin D1 (2926), pRB Ser780 (9307), p-4EBP1 Ser65 (9451), Thr70

(5078) and Thr37/46 (2855) and 4EBP1 (9452) (Cell Signaling Technologies) and GAPDH (8245) (Abcam, Cambridge, Massachusetts). Secondary anti-mouse and anti-rabbit immunoglobulin conjugated with horseradish peroxidase (Santa

Cruz Biotechnology, Dallas, Texas) were used at a 1:3000 dilution in 5% milk/TBS-T. The blots were visualized with ECL Western Blotting Detection

Reagent (Thermo, Waltham, Massachusetts) using the ChemiDoc XRS+ Imager

(Bio-Rad).

4.2.5 In Vivo Xenograft Study 1 x 107 ACHN cells in matrigel and sterile saline (0.1mL) were injected into the right flank of 5 - 7 week old male SCID mice. The animals were housed according to a 12:12 hr light-dark cycle and allowed food and water ad libitum.

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When the mean tumor volume reached approximately 500 mm3 (approximately day 16), mice were pair-matched into the following 4 groups: pentoxifylline vehicle + sorafenib vehicle, sorafenib (10 mg/kg) + pentoxifylline vehicle, sorafenib vehicle + pentoxifylline (40 mg/kg), and sorafenib (10 mg/kg) + pentoxifylline (40 mg/kg), (n = 4). Sorafenib was dosed once daily via oral gavage with Cremephor EL/ethanol/water (12.5 : 12.5 : 75) solution and pentoxifylline was administered once daily intraperitoneally (ip) in PBS for a period of 14 days. Tumor volume was estimated approximately twice weekly using the formula v = ½ x a x b2, where v is the volume (mm3), a is the long diameter (mm) and b is the short diameter (mm). Tumor growth inhibition (TGI) was calculated using the formula TGI = [1 - (T/C) x 100], where T and C are the mean tumor volume in the treated and untreated control groups, respectively. All procedures were completed in accordance with the University of Arizona

Institutional Animal Care and Use Committee.

4.2.6 Xenograft Tissue Harvest and Analysis Preparation

At the completion of the study, mice were euthanized via CO2 asphyxiation

2 hrs following final drug dose. Pictures of the mice were taken and subcutaneous xenograft tissue was harvested and divided in half. One half was frozen in N2(l) for tissue lysate generation followed by Western blot analysis. The other half was fixed for immunohistochemistry (IHC) analysis following the neutral buffered formalin (NBF; 10%) 2 x 2 method (Chafin et al., 2013). Briefly, xenograft tissue was submerged in chilled NBF and stored at 4ºC for 2 hrs, then

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placed in a 45ºC warm box for 2 hrs and finally transferred to 70% ethanol (v/v) at room temperature prior to paraffin embedding.

4.2.7 Immunohistochemistry Paraffin embedded blocks were cut into 4-micron sections.

Immunohistochemistry assays were performed on a VENTANA Discovery XT automated staining instrument according to the manufacturer’s instructions

(Ventana Medical Systems, Tucson, Arizona). Slides were de-paraffinized using

EZprep solution (Ventana Medical Systems) at 90 ºC, and all reagents and incubation times were chosen as directed on antibody package inserts. Slides were developed using the OmniMap DAB detection kit (Ventana Medical

Systems) and counterstained with hematoxylin. Antibody titers were determined for each antibody using positive and negative control tissues following the manufacturer’s instructions.

Histologic assessment was scored using a modification of a scoring system (modified H-score) published by the manufacturer of the tissue processor

(Leica Biosystems, Buffalo Grove, Illinois) and further described in a manuscript detailing the 2 x 2 fixation method (Chafin et al., 2013). Briefly, a blinded pathologist graded each tissue section from 0 to 3+ (with 3+ being highest) in regions pertaining to the nuclear, cytoplasmic and overall morphology, based on the intensity of staining. IHC H-score was calculated using the following equation,

H-Score =

[1 x (% cells @ 1+) + 2 x (% cells @ 2+) + 3 x (% cells @ 3+)]. Thus, and

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individual H-score can range from 0 to 300, with 300 reflecting 100% of the cells displaying 3+ staining. Differences in histologic quality scores were assessed with the Kruskal-Wallis test and Dunn’s post-test to correct for multiple comparisons.

4.2.8 Statistics Data are expressed as mean averages ± standard error, unless otherwise stated. Mean values were compared using a Student's T-test, two tails, with equal variances. P-values are expressed as *p<0.05, **p<0.01, and ***p<0.001.

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4.3 Results

4.3.1 Antiproliferative effects of sorafenib in ACHN are time- and dose- dependent

In order to explore PTXs use as an adjuvant therapy in RCC treatment, we investigated whether PTX potentiated the antiproliferative effects of sorafenib in

ACHN cells. To determine the optimal dosage of sorafenib, ACHN cells were treated with a dose range from 5 to 50 µM of sorafenib (similar to doses used in

QTRRE studies performed previously in our lab) for 2 hrs (Figure 4.1A), and mitochondrial dehydrogenase activity was measured using the MTS assay. It was determined that 50 µM sorafenib at 2 hrs provided an optimal response, with a decrease of relative mitochondrial dehydrogenase activity to 57.78 (SD ±

5.03%, p<0.01) of untreated control. A time course was also conducted with 25

µM sorafenib from 0 to 6 hrs (Figure 4.1B). Treatment with 25 µM sorafenib at 2,

4, and 6 hrs caused a decrease of relative mitochondrial dehydrogenase activity to 77.83 (SD ± 6.61%, p<0.05), 69.98 (SD ± 4.02%, p<0.01), and 65.74 (SD ±

4.20%, p<0.01) compared to untreated control, respectively.

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Figure 4.1 Antiproliferative effects of sorafenib in ACHN cells

ACHN cells were treated with sorafenib [soraf] for 2 hrs at the concentrations shown (A) or at 25 µM for the durations shown (B). Mitochondrial dehydrogenase activity was analyzed using the CellTiter 96® Aqueous Non-Radioactive Cell Proliferation (MTS) Assay. Values are means and error bars show SD (n=3). P- values are expressed as *p<0.05, **p<0.001, compared to untreated control.

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4.3.2 Combination of PTX + sorafenib treatment significantly decreases cyclin D1 and proliferation in ACHN cells

ACHN cells were treated with PTX (1 mg/ml) and sorafenib (50 uM) alone, and in combination for 2 hrs to determine whether PTX potentiated sorafenib in regards to decreasing cyclin D1 levels. Analyzing cyclin D1 levels via Western blot demonstrated that combination treatment enhanced cyclin D1 decrease over each drug alone (Figure 4.2A). Further, ACHN cells were pre-treated with PTX

(1.0 mg/mL) for 24 hrs alone, sorafenib (50 µM) for 4hrs alone, or combination of

PTX pre-treatment followed by sorafenib treatment in an effort to demonstrate

PTXs ability to potentiate the antiproliferative effects of sorafenib. Combination treatment with PTX and sorafenib displayed enhanced antiproliferative effects in

ACHN over each drug alone (Figure 4.2B). Sorafenib alone caused a 66.86 (SD

± 3.81%, p<0.05) fold decrease of relative mitochondrial dehydrogenase activity, whereas combination treatment caused a decreased fold of 47.31 (SD ± 4.57%, p<0.01), compared to untreated control.

Finally, ACHN cells were again treated with various doses of PTX alone for 24 hrs, various doses of sorafenib alone for 2 hrs, and a combination of both drugs. MTS analysis shows that pre-treatment of PTX (1 mg/ml) for 24 hrs, followed by 5 µM sorafenib for 2 hrs achieves greater antiproliferative effects than 50 µM sorafenib alone at 2 hrs (Figure 4.3). PTX, presumably through G1

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phase arrest, was capable of dose-lowering the effective sorafenib treatment from 50 to 5 µM in ACHN cells, a desirable effect for a possible adjuvant treatment. These results led us to investigate whether PTX conferred similar adjuvant properties in in vivo models using ACHN xenografts in mice.

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Figure 4.2 Increased antiproliferative efficacy with PTX + sorafenib treatment in ACHN cells (A) Whole cell lysates were generated from ACHN cells treated with PTX (1.0 mg/mL), soraf (50µM), or a combination of PTX + soraf for 2 hrs. Equal amounts of protein were separated via SDS-PAGE and immunoblotted for cyclin D1 and GAPDH. (B) ACHN cells were pre-treated with PTX (1.0 mg/mL) for 24 hrs, soraf (50µM) for 4hrs, or combination of pre-treatment followed by soraf treatment. Proliferation was measured via mitochondrial dehydrogenase activity using the CellTiter 96® Aqueous Non-Radioactive Cell Proliferation (MTS) Assay. Values are means and error bars show SD (n=3). P-values are expressed as *p<0.05, **p<0.01, compared to untreated control.

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Figure 4.3 PTX pretreatment lowers the effective dose of sorafenib in ACHN cells ACHN cells were pre-treated with PTX for 24 hrs with the concentrations shown, soraf for 2 hrs with the concentrations shown, or combination of pre-treatment followed by soraf treatment as shown. Proliferation was measured via mitochondrial dehydrogenase activity using the CellTiter 96® Aqueous Non- Radioactive Cell Proliferation (MTS) Assay. Values are means and error bars show SD (n=5). P-values are expressed as **p<0.01, ***p<0.001.

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4.3.3 Combination of PTX + sorafenib treatment inhibits ACHN tumor growth

In order to test the efficacy of PTX and sorafenib in vivo, we utilized the

ACHN xenograft model. Mice bearing tumors were divided into 4 groups: pentoxifylline vehicle + sorafenib vehicle [veh], sorafenib (10 mg/kg) + pentoxifylline vehicle [soraf], sorafenib vehicle + pentoxifylline (40 mg/kg) [PTX], and sorafenib (10 mg/kg) + pentoxifylline (40 mg/kg) [combo]. Following 14 days of treatment, all treatment groups displayed tumor growth inhibition, compared to control (Figure 4.4). Tumor volume differences were observed in mice receiving different therapies, as shown in representative mouse images (Figure 4.4A). At the termination of the study, vehicle mean tumor volume reached 1203 (SE ± 43 mm3), soraf mean tumor volume reached 963 (SE ± 138 mm3), PTX mean tumor volumes reached 1097 (SE ± 111 mm3), and combination mean tumor volume reached 787 (SE ± 68 mm3) (Figure 4.4B). This corresponds to a tumor growth inhibition (TGI) of 19.93%, 8.78%, and 34.57% for soraf, PTX, and combo, respectively. Only combination dose demonstrated significant TGI and fold decrease in mean tumor volume (p<0.05). No therapy-related toxicity was observed, demonstrated by equal mouse bodyweights (Figure 4.4C).

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Figure 4.4 In vivo ACHN tumor growth inhibition via PTX + sorafenib treatment When mean tumor volumes reached approximately 500 mm3, mice were pair- matched into the following 4 groups: pentoxifylline vehicle + sorafenib vehicle [veh], sorafenib (10 mg/kg) + pentoxifylline vehicle [soraf], sorafenib vehicle + pentoxifylline (40 mg/kg) [PTX], and sorafenib (10 mg/kg) + pentoxifylline (40 mg/kg) [combo], treated for 14 days. A. Representative images of mice bearing tumors dosed with the therapies shown above. B. Mean tumor volumes (mm3) in mice dosed with therapies shown. C. Mean mouse bodyweights (g) in mice dosed with therapies shown. Values are means (n = 4). P-values are expressed as *p<0.05.

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4.3.4 Combination of PTX + sorafenib treatment decreases pRb, p4EBP1, and cyclin D1 levels in ACHN xenograft tissue

We wished to validate whether combination treatment with PTX and sorafenib caused significant decreases in cyclin D1 in ACHN xenograft tissue, concomitant with TGI, as previously observed in ACHN cells (Figure 4.2). In addition, we investigated relative pRb, and p4EBP1 levels in ACHN xenograft tissue, following treatment for 14 days. Via Western blot analysis, we observed decreases in pRb, p4EBP1 Ser 65 and Thr 70, as well as cyclin D1 levels in PTX and sorafenib treated xenografts, both alone and in combination (Figure 4.5).

Further, combination treatment caused significant decrease in cyclin D1 levels, compared to either drug treated alone in the ACHN xenograft tissue (p<0.05), concomitant with increased TGI. Of note, PTX alone induced similar levels of decreased pRb, p4EBP1 Ser 65, Thr 70, and cyclin D1 seen in sorafenib alone treated xenografts. Further, PTX alone promoted loss of p4EBP1 Thr 37/46, compared to no change observed following sorafenib alone treatment. These results are the first to demonstrate PTX treatment caused decreased p4EBP1 levels, concomitant with cyclin D1 levels, in vivo.

Via IHC analysis, we validated a decrease in cyclin D1 levels observed in combination treated xenograft tissue (Figure 4.6). Combination treatment promoted a decreased modified H-score for cyclin D1 protein levels, compared to either drug treated alone, though significance of p<0.5 was not achieved.

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Figure 4.5 PTX + sorafenib decrease pRB, p4EBP1, and cyclin D1 levels in ACHN xenograft tissue, via Western blot analysis

Following 14 days of treatment with vehicle [V], sorafenib (10 mg/kg) [S], PTX (40 mg/kg) [P], or a combination of PTX + sorafenib [C], ACHN xenograft tissue was harvested and frozen in N2(l) for tissue lysate generation prior to Western blot analysis for the targets shown above. Image shows representative figure. Values are means, bars show SE, (n = 4). P-values are expressed as *p<0.05, compared to untreated control.

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Figure 4.6 PTX + sorafenib decrease cyclin D1 levels in ACHN xenograft tissue, via IHC analysis Following 14 days of treatment with vehicle [V], sorafenib (10 mg/kg) [S], PTX (40 mg/kg) [P], or a combination of PTX + sorafenib [C], ACHN xenograft tissue was fixed for immunohistochemistry (IHC) analysis following the neutral buffered formalin (NBF; 10%) 2 x 2 method (see methods section for additional info). IHC was carried out for cyclin D1 protein levels. Values are means, bars show SD, (n = 4).

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4.4 Discussion

In this study we validated PTX drug effects, previously observed in ACHN cells (Chapter 3), in xenograft tissue. We observed that treatment with PTX caused decreased p4EBP1 Ser65, Thr 70, and Thr 37/46 levels in xenograft tissue that were concomitant with decreased cyclin D1 and pRb levels (Figure

4.5), validating our proposed mechanism of action behind cyclin D1 decrease.

These represent novel drug effects of PTX in cancerous tissue. Interestingly,

PTX was just as effective as sorafenib in decreasing p4EBP1, cyclin D1, and pRB levels in these xenografts. We also observed PTX to yield additive effects, with regard to cyclin D1 decrease and inhibition of cell proliferation in ACHN cells when dosed in combination with sorafenib (Figures 4.2 and 4.3). With this knowledge, we decided to test the efficacy of PTX and sorafenib in an in vivo

ACHN xenograft model. Consistent with our findings in cells, PTX and sorafenib treatment caused significant decrease in cyclin D1 as well as significant tumor growth inhibition, compared to either drug when treated alone (Figure 4.4 and

4.5).

Our findings agree with previous reports on PTX tumor growth inhibition in xenografts models. PTX (40 and 60 mg/kg) inhibited A375 melanoma tumor growth by 35 and 50% in xenograft models, respectively (Kamran and Gude,

2012). In addition, PTX (40 and 60 mg/kg) were found to inhibit MDA-MB-231 tumor growth, equally, by 36% in xenograft models (Goel and Gude, 2013).

Further, combination treatment with PTX and doxorubicin (DOX) delayed tumor 151

growth more than either treatment alone in breast cancer xenograft models (Goel and Gude, 2014b). Investigation into cyclin D1 levels or perturbation in upstream signaling via PTX was not conducted in these tissues. Our findings that PTX decreased p4EBP1 and cyclin D1 levels (Figure 4.5) add evidence to possible mechanisms regarding the antitumor effects of PTX in these previously observed studies. These effects mimic the action of the FDA approved RCC targeted therapies, everolimus and temsirolimus, that inhibit mTOR, effectively reducing

4EBP1 phosphorylation and subsequent translation of proteins, like cyclin D1, believed to contribute to cancer progression (Faivre et al., 2006; Jastrzebski et al., 2007), and further support its use as an adjuvant treatment in treating RCC.

PTX was also found to decrease elevated tumor interstitial fluid pressure

(TIFP) as well as increase pO2 in murine fibrosarcoma tumors, indicative of increased tumor perfusion (Lee et al. 2000). These findings demonstrate a significant ability of PTX in increasing oxygenation, systemic drug delivery, and ameliorating hypoxia in solid tumors, presumably through its hemorheologic effects. In addition to increasing tumor perfusion, PTX was discovered to increase intracellular concentrations of doxorubicin (DOX) and circumvent resistance in multidrug resistant (MDR) P388/DOX leukemia cells (Viladkar and

Chitnis, 1994) as well as vincristine (VCR) in MDR L1210/VCR mouse leukemic cells (Breier et al., 1994), via a significant downregulation of Pgp in these models. Whether these mechanisms are partly responsible for the synergistic

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effect observed between PTX and sorafenib in ACHN cell and xenograft growth inhibition (Figures 4.2B, 4.3 and 4.4B) is unknown.

Sorafenib is highly cell-permeable and considered a weak Pgp substrate in vitro (Dagenais et al., 2009), suggesting possible decreased Pgp expression mediated by PTX is not a factor in the increased antiproliferative capacity of combination treatment in ACHN cells or xenografts. In addition, cyclin D1 levels, consistent with cell and tumor growth inhibition, were found significantly decreased in combination treatment with PTX and sorafenib (Figures 4.2 and

4.5), compared to either treatment alone, suggesting G1 phase cell cycle arrest is a factor in the observed synergy between these drugs. Further research is needed to confirm whether PTXs effects on pAKT/p4EBP1/cyclin D1 modulation are solely responsible for its observed additive antitumor effects. Regardless, results from this study strengthen the possible adjuvant role that PTX may serve, alongside sorafenib and other targeted therapies, in treating RCC.

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CHAPTER 5: CONCLUDING REMARKS

5.1 Antiproliferative effects of PTX in multiple RCC cancer models

QTRRE, a rodent renal epithelial cell line transformed by the active metabolite of hydroquinone, TGHQ, are null for the tumor suppressor, tuberin, encoded by the TSC-2 gene (Yoon et al. 2001). QTRRE cells are tumorigenic in athymic nude mice and produce tumors that resemble human RCC cells (Patel et al., 2003). The ACHN human RCC cell model are null for VHL, reflective of RCC tumors found in humans (Gnarra et al. 1996; Walker et al. 1998). Human VHL

RCC and rodent Tsc-2 RCC not only share clear cell pathology, but also a number of common downstream targets. VHL and TSC-2 null RCC models both result in accumulation of HIFα leading to increased protein levels of VEGF, as well as activation of MAPK and PI3K/AKT/mTOR/4EBP1 signaling, leading to overexpression of cyclin D1 and enhanced cell proliferation rates (Brugarolas et al., 2003; Liu et al., 2003; Rebuzzi et al., 2007). We found that PTX induced cyclin D1 decrease in QTRRE, and ACHN, as well as human RCC Caki-2, and

786-O RCC cell models, and significantly arrested the QTRRE, ACHN, and Caki-

2 models in the G1 phase (Chapter 2).

These findings are consistent with observations made in melanoma cells

(Kamran & Gude, 2012; Sztiller-Sikorska et al., 2014), T-cell lymphoma cells

(Rishi et al., 2009), B16F10 melanoma cells (Dua and Gude, 2006), MDA-MB-

231 breast cancer cells (Goel and Gude, 2011) and U937 leukemia cells (Bravo- 154

Cuellar et al., 2013). Ongoing studies in our laboratory effectively demonstrate that PTX significantly decrease cyclin D1 levels in PC3, DU145, and LNCaP human prostate cancer cell models as well. These effects were both PTX dose- and duration-dependent. Together, these results suggest that PTX may be effective in inhibiting the growth of vastly different cancer types characterized by cyclin D1 overexpression.

Despite observing cyclin D1 decrease across all RCC models tested, the human 786-O human RCC cell line exhibited insignificant G1 phase arrest following 24 hr PTX treatment (Figure 2.4). 786-O also displayed rapidly recovered expression of cyclin D1 levels (Figure 2.6) that was not due to increased transcription of the CCND1 gene (Figure 3.1E). Mechanism(s) behind such rapid recovery in 786-O are currently unknown. Disparate responses to

PTX amongst these RCC cell models represents an example of a need for personalized medicine. Through further understanding of the respective tumorigenic drivers responsible for growth, better therapeutic regimens can be utilized on a patient to patient basis.

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5.2 Targeting pAKT/p4EBP1/cyclin D1 in RCC

Our results showed PTX promoted pAKT inhibition (Figure 3.2) which is consistent with previously published studies. PTX inhibited the PDGF-activated phosphorylation of AKT in rat airway smooth muscle cells (Chiou et al., 2006) as well as AKT activation in fMLP-stimulated human neuronal cells (Costantini et al.,

(Figure 3.3) and xenografts (Figure 4.5). These results represent novel PTX drug effects, and correlate decreased pAKT with decreased cyclin D1 protein levels, as previously observed (Lin et al., 2003).

Cyclin D1 overexpression is a common event in cancer but does not occur solely as a consequence of gene amplification. Rather, increased levels of cyclin

D1 frequently result from its defective regulation at the post-translational level, primarily in processes involved with Thr 286 phosphorylation, nuclear export, and ubiquitin-mediated proteasomal degradation (Gillett et al., 1994; Russell et al.,

1999). Indeed, several mutations that impair phosphorylation-dependent nuclear export of cyclin D1 have been identified in endometrial cancer (Ikeda et al., 2013;

Moreno-Bueno et al., 2003) as well as in esophageal carcinoma and esophageal cancer-derived cell lines (Benzeno et al., 2006). Further, high levels of cyclin D1b

(lacking the C-terminal Thr286 residue) have been reported to be associated with

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a poor prognosis for breast cancer patients (Abramson et al., 2010; Asiedu et al.,

2014; Taneja et al., 2010; Zhu et al., 2010) as well as non-small cell lung cancer patients (Li et al., 2008). Because PTX appears to inhibit the translation of cyclin

ACHN (Figure 3.8C).

5.3 PTX and combinatorial therapeutics in RCC

The basic principles behind combination therapy are to overcome drug resistance, decrease toxicity and/or improve efficacy (Zimmermann et al., 2007).

With respect to toxicity profile, sorafenib is not alone amongst the FDA approved

RCC treatments. All of the FDA approved targeted RCC therapies inflict commonly observed moderate to severe adverse effects. Combinations of these drugs are currently undergoing extensive research in hopes decreasing the effective dose of each respective drug, thus lowering the chance of adverse reactions. Currently, it is difficult to find an ongoing clinical study using only one of the targeted drugs given in monotherapy. In addition to lowering toxicity, combination therapy has the potential to raise efficacy and overcome resistance that is often seen in tumors when treated with a single agent.

Unfortunately, to date, no effective adjuvant treatment for RCC has been described (including radio-, immuno- and chemotherapies), though numerous 157

trials involving targeted agents have not yet reported results (Janowitz et al.,

2013). With a 5-year relapse rate for intermediate- and high-risk early-stage RCC of 30-40% (Escudier et al., 2012), research in the area of combination and adjuvant therapies for RCC is crucial. Advances in our understanding of the molecular pathogenesis of RCC will identify additional potential targets for adjuvant treatment, which should expand and diversify adjuvant treatment options.

We found that PTX increased the antiproliferative capacity of sorafenib in

ACHN cells (Figures 4.2 and 4.3) as well as in ACHN xenografts (Figure 4.4B), presumably through inducing cell cycle arrest in the G1 phase. PTX serves as an attractive therapeutic because it is well tolerated. Less than 10% of patients experience adverse effects such as dizziness, nausea, and indigestion. Less that

1% experience more serious adverse effects such as angina and palpitations.

PTX also has minor to insignificant drug interactions with other hemorheologic therapeutics (e.g. warfarin). Further, PTXs palliative effects (Bese et al., 2007;

Hille et al., 2005; Kim and Park, 2006; Mantovani and Madeddu, 2010; Melo et al., 2008; Steeves and Robins, 1998; Tazi and Errihani, 2010; Theis et al., 2010;

Worthington et al., 2010), as well as previously observed tumor sensitizing effects (Breier et al., 1994; Drobná et al., 2002; Krishna and Mayer, 2000;

Kvackajová-Kisucká et al., 2001; Nieder et al., 2005; Viladkar and Chitnis, 1994), and recently documented antitumor effects (Bravo-Cuellar et al., 2013; Goel and

Gude, 2014a, 2014b, 2013; Kamran and Gude, 2013, 2012) strengthen the idea for its use as an adjuvant in the treatment of RCC. 158

CHAPTER 6: FUTURE DIRECTIONS

6.1 Overview

Our data demonstrate the novel ability of PTX to arrest cancer cells in the

G1 phase via a decrease in p4EBP1, cyclin D1, and pRb levels. It is known that

PTX is a non-specific PDE inhibitor. What is not currently known is whether such inhibition leads to the observed cyclin D1 decrease and G1 phase cell cycle arrest. Thus, PTXs target, with respect to these drug effects, is currently unknown. Identification of the initial target will be important, should a more potent, cyclin D1 ablative therapeutic be desired.

Parallel to target identification, drug structure optimization is of interest. Of the methylxanthine derivatives tested in these studies (PTX, theophylline, IBMX and db-cAMP), only PTX conferred cyclin D1 decrease. Differences in pharmacologic effect are dependent on the inherent structure of the therapeutic and understanding how specific structure alterations to PTX affect cyclin D1 decrease may help lead to an optimized, more potent drug.

Cancer is a highly personalized disease and, thus, will require highly personalized treatment regimens. Our human RCC cell models displayed varying sensitivity to PTX with respect to both cyclin D1 decrease and G1 phase cell cycle arrest. Understanding inherent differences in cyclin D1 regulation between these models may assist in identifying possible predictive biomarkers for PTX response. Identification of the initial target discussed above may help in

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identifying predictive biomarkers for PTX response. Alternatively, biomarker identification may help in identifying PTXs initial target.

Finally, these studies are the first to demonstrate that PTX has an anticancer role, both alone and in combination with sorafenib, in ACHN cells and xenografts. These data are parallel with the few recent studies demonstrating

PTX to have anti-cancer properties in a number of other cancer types (Goel and

Gude, 2014a, 2013, 2011; Kamran and Gude, 2012). Understanding which cancer types respond to PTX will help nominate its use as a co-therapy. Further, understanding its utility as an adjuvant therapy alongside other, targeted anticancer therapeutics is desired.

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6.2 PTX target identification

6.2.1 Introduction

Our data demonstrate that PTX is able to arrest cancer cells in the G1 phase via a decrease in cyclin D1 and subsequently pRb levels (Chapter 2).

CCND1 transcription or cyclin D1 protein stability do not appear to be affected by

PTX treatment. Cyclin D1 decrease may occur through a novel mechanism of

PTX by which 4EBP1 phosphorylation decreases, resulting in decreased translation (Chapter 3 and 4). 4EBP1 phosphorylation can be modulated via a number of upstream kinase pathways including the PI3K/AKT/mTOR pathway as well as the Raf-1/MEK/ERK MAPK pathway (Cohen et al., 2011; Pópulo et al.,

2012). Exploration of the activation states of these upstream kinase pathways may yield information on the initial target of PTX. Understanding the mechanism(s) responsible for p4EBP1 loss may help to further describe novel properties of PTX.

In addition, knowledge that PTX, but not db-cAMP nor theophylline, caused cyclin D1 decrease (Figure 2.1) may shed light on the initial drug target.

Because PTX is a non-specific PDE inhibitor, it is not know which of the 11 PDE families may be responsible for PTXs decrease in cyclin D1. Using PDE-specific inhibitors to replicate cyclin D1 decrease may identify particular PDE(s) that are responsible for this drug effect. In addition, genetic PDE knock-out studies would validate which PDEs are responsible for cyclin D1 decrease and G1 phase

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arrest. Such a target may be amenable to more potent small molecule intervention for the adjuvant treatment of RCC and other cancers.

6.2.2 Methods

A. Detection of upstream p4EBP1 modulator activity via Western blot and in-vitro kinase assay

Whole cell lysates following PTX treatment will be subjected to 10, 12 or

14% SDS-PAGE at 140V, followed by electrophoretic transfer to PVDF membranes. Western blot primary antibodies to be used are p-4EBP1 Ser65

(9451), Thr70 (5078) and Thr37/46 (2855) and 4EBP1 (9452), p-mTOR Ser2448

(2971) and Ser2481 (2974), pAKT Ser473 (9271) and Ser308 (9275), pPI3K

Tyr458/Tyr199 (4228), p-p44/42 MAPK (Erk1/2) Thr202/Tyr204 (9101) (Cell

Signaling Technologies) and GAPDH (8245) (Abcam). Upstream proteins displaying decreased activity following PTX treatment will be immunoprecipitated using protein specific antibodies bound to protein A/G-agarose beads (Pierce

Biotechnology). Kinase activity of the IPd kinases will be determined using the

Cascade Assay kits (Upstate Biotechnology) as previously reported (Yoon et al.,

2004). The incorporated [γ-33P] ATP will be quantified via liquid scintillation spectroscopy.

B. Detection of possible PDE target via PDE-specific small molecule inhibitors and PDE-specific siRNA studies

Whole cell lysates following known PDE-specific inhibitor treatment (PDE:

-1; Vinpocetine, -2; erythro-9-(2-hydroxy-3-nonyl)adenine [EHNA], -3; cilostazol, -

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4; rolipram, -5; sildenafil/tadalafil, -7; quinazoline, -10; papaverine) will be generated. In addition, SMARTpool ON-TARGETplus PDE 1-11 and siGENOME non-targeting control.siRNA (Dharmacon RNAi Technologies, Lafayette,

Colorado) will be transfected into cells using DharmaFECT 1 Transfection

Reagent (Dharmacon), according to manufacturer's protocol. These lysates will be subjected to 10, 12 or 14% SDS-PAGE at 140V, followed by electrophoretic transfer to PVDF membranes. Western blot primary antibodies to be used are cyclin D1 (2926) (Cell Signaling Technologies) and GAPDH (8245) (Abcam).

6.2.3 Anticipated results Following PTX treatment, Western blot analysis and in-vitro kinase assay analysis may yield decreased activity of candidate upstream regulators of 4EBP1 phosphorylation (mTOR/ERK). These findings would help explain the loss of phosphorylation observed on 4EBP1 following PTX treatment, and would further describe the novel drug effect. We anticipate such a decrease in either or both of these upstream pathways may be occurring. With knowledge of specific upstream signaling modulation, identification of PTXs target may be clearer.

PDEs can either convert cGMP to GMP or cAMP to AMP or both. The known pharmacologic actions of PTX stem from non-specific inhibition of PDEs, particularly those that convert cAMP to AMP (PDE 4, 7, and 8). Should PTXs decrease in cyclin D1 and induction of G1 phase arrest be through inhibition of one or a combination of these, we anticipate treatment with selective inhibitors

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and/or siRNA knockdown of PDEs 4, 7, and 8 to replicate such drug effect. This would help suggest whether PDEs are the target for PTX-mediated cyclin D1 decrease and G1 phase arrest.

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6.3 Drug structure optimization

6.3.1 Introduction

Of the methylxanthine derivatives tested in QTRRE and ACHN cells, only

PTX demonstrated cyclin D1 decrease. In a similar fashion to our observations,

PTX (but not xanthines: theophylline or caffeine) reversed vincristine resistance of drug-resistant L1210/VCR mouse leukemic cells (Stefanková et al., 1996). The authors of this study concluded that reversal of vincristine resistance could not be explained from the point of known pharmacological effects of PTX that are common for other xanthines such as inhibition of PDE activity. In agreement, it was discovered that PTX induces the sensitization of the above cells to vincristine via downregulation of Pgp (Drobná et al., 2002), a novel drug effect at the time. Considering disparities in the structure and efficacy of the xanthines previously tested, 25 analogues of PTX (composed of different substituents located in positions N1, N3, N7 and C8) were tested in combination with vincristine with respect to cell survival, [3H]-vincristine accumulation, and Pgp mRNA expression in L1210/VCR cells (Kupsáková et al., 2004). The authors concluded that existence of a long, polar substituent at the N1 position

(characteristic of PTX [Figure 1.8], but not theophylline, IBMX or db-cAMP) plays a crucial role in the effective reversal of Pgp-mediated multidrug resistance in the above cells. Further, they concluded that elongation of the substituent at positions N3 and N7 (from methyl to propyl) increases and at the position C8

(from H to propyl) decreases the efficacy of xanthines to reverse the vincristine 165

resistance of L1210/VCR cells. Further, this group synthesized a new PTX derivative, 1-(10-undecylenyl)-3-heptyl-7-methyl xanthine (PTX-UHM), with a prolonged substituent at the N1 position and found it to be significantly more efficient in reversing VCR resistance in the L1210/VCR cell line, compared to

PTX (Docolomanský et al., 2005).

Parallel to these studies, it may be advantageous to test the efficacy of

PTX analogues with respect to cyclin D1 decrease and G1 phase arrest in

ACHN, in order to develop a more potent therapeutic.

6.3.2 Methods

A. Western blot analysis of cyclin D1 in ACHN treated with differing PTX analogues

Whole cell lysates of ACHN following treatment with PTX and various PTX analogues (including PTX-UHM) will be subjected to 10, 12 or 14% SDS-PAGE at 140V, followed by electrophoretic transfer to PVDF membranes. Western blot primary antibodies to be used are cyclin D1 (2926) (Cell Signaling Technologies) and GAPDH (8245) (Abcam).

B. Detection of cell cycle arrest in ACHN treated with differing PTX analogues

PTX analogues yielding enhanced cyclin D1 decrease in ACHN should be tested via cell cycle arrest to validate increased efficacy in inducing G1 phase arrest, compared to PTX. Following specific drug combination treatment, cells will be fixed in 70% EtOH. 25 µl (1/20 vol) of 10 mg/mL RNAse A and 12.5 µl (1/40

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vol) of 1.6 mg/mL propidium iodide will be added into each tube. Tubes will incubate at 37 ºC for 30 min, and undergo analysis via flow cytometry.

6.3.3 Anticipated Results

The ultimate drug target behind PTX-mediated downregulation of Pgp is still unknown. Depending on the initial drug target of PTX, with respect to cyclin

D1 decrease and G1 phase arrest, specific PTX analogs may yield higher efficacy. Should the target behind Pgp decrease and cyclin D1 decrease be the same, we would expect to find similar results to the studies mentioned previously. Because PTX-mediated decrease of Pgp was dependent on a decrease in transcription of Pgp mRNA, but not cyclin D1 mRNA, we suspect the ultimate drug target to differ. Nonetheless, because PTX, but not other methylxanthines, showed this novel effect, drug optimization still represents an important goal.

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6.4 Biomarker identification for predicting PTX response/sensitivity

6.4.1 Introduction

Signal transduction pathways regulating cyclin D1 protein levels are affected by complex synthesis (transcription and translation) and degradation processes. Our cell models displayed varying sensitivity to PTX with respect to decrease of cyclin D1 protein levels as well as G1-phase arrest (Chapter 2).

Understanding the inherent differences between our cell models with regard to cyclin D1 regulation may help explain the heterogeneity seen in PTX sensitivity.

Further, identification of predictive PTX-sensitivity biomarker(s) is desired.

6.4.2 Methods

A. Western blot analysis of candidate biomarkers related to cyclin D1 regulation in PTX-sensitive and PTX-resistant cell models

Western blot analysis of candidate biomarkers related to cyclin D1 regulation in PTX-sensitive and PTX-resistant cell models may help in identifying biomarkers capable of predicting PTX response. Comparing relative levels of candidate biomarkers between these PTX-sensitive and -resistant cells lines is one of the easiest ways of identifying such biomarkers. Possible targets to investigate would be well-studied proteins that have a known effect on cyclin D1 synthesis, as it appears this is the step with which PTX interferes to decrease overall levels. Our data suggests that modulation of the pAKT/p4EBP1 pathway is responsible for PTXs decrease of cyclin D1. Comparing relative expression

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levels of proteins within and related to this pathway via Western blot analysis is a logical first approach.

B. Candidate biomarker validation via transfection and siRNA

Candidate biomarkers identified in PTX-sensitive models (QTRRE,

ACHN), can be transfected into PTX-resistant models (786-O), if they were found to be originally decreased or lacking in expression. As an example, p27 was found to be highly expressed in QTRRE and ACHN, however, 786-O revealed very low levels of p27 levels (Figure 2.5C). Upregulation of p27 via transient or stable transfection may rescue PTXs ability to decrease cyclin D1 and induce G1 phase arrest in 786-O, consistent with our findings in the QTRRE and ACHN cell lines. Such a rescue would validate high p27 levels as being predictive in PTX- sensitivity. Likewise, candidate biomarkers identified in PTX-resistant models, but lacking in PTX-sensitive models, can be downregulated via siRNA silencing. As an example, tuberin, the TSC-2 gene product, was found to be highly expressed in 786-O but low in ACHN and QTRRE. Following knockdown of tuberin in 786-

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6.5 PTX adjuvant activity alone and in combination with other targeted, anti-cancer therapeutics in ACHN and other cancer cell types

6.5.1 Introduction Our data demonstrates that PTX has both a dose-lowering as well an additive effect in decreasing proliferation in vitro with the FDA-approved BRAF and VEGF inhibitor, sorafenib, in ACHN cells (Chapter 4). PTX also had an additive effect alongside sorafenib on tumor growth inhibition of ACHN xenograft tumors (Chapter 4). Presumably, other targeted therapeutics may benefit from the G1 cell cycle arrest caused by PTX. Testing pre-treatment of PTX with other

FDA-approved RCC therapies in ACHN cells represents a logical start. Further, the utility of PTX dosed alone and in combination with other anti-cancer therapies in different cancer cell types (particularly those marked by cyclin D1 overexpression; NSCLC; melanoma; breast; pancreatic; prostate) should be explored. Recent results from our laboratory definitively reveal that PTX significantly decreases cyclin D1 in LNCaP, DU145 and PC3 human prostate cancer cells. Concomitant with cyclin D1, PTX also decreases pAKT (Ser 308 and Ser 473) as well as p4EBP1 levels in LNCaP cells. Further investigation is ongoing at the time of writing this dissertation. These results show promise that

PTX may yield such effects in other cancer cell types.

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6.5.2 Methods

A. Western blot analysis of p4EBP1, cyclin D1 and pRb in different drug combination-treated cells Whole cell lysates as well as cytosolic/nuclear protein fractions following specific drug combination treatment will be subjected to 10, 12 or 14% SDS-

PAGE at 140V, followed by electrophoretic transfer to PVDF membranes.

Western blot primary antibodies to be used are p-4EBP1 Ser65 (9451), Thr70

(5078) and Thr37/46 (2855) and 4EBP1 (9452), cyclin D1 (2926), pRB Ser780

(9307), SP1 (5931) (Cell Signaling Technologies) and GAPDH (8245) (Abcam).

B. Detection of cell cycle arrest

Drug combinations yielding enhanced p4EBP1, cyclin D1 and pRb decrease in ACHN and other cancer types should undergo cell cycle analysis to further validate the adjuvant utility of PTX. Following specific drug combination treatment, cells will be fixed in 70% EtOH. 25 µl (1/20 vol) of 10 mg/mL RNAse A and 12.5 µl (1/40 vol) of 1.6 mg/mL propidium iodide will be added into each tube. Tubes will incubate at 37 ºC for 30 min, and undergo analysis via flow cytometry.

C. Detection of apoptosis

Drug combinations yielding enhanced sub-G1 peaks via cell cycle analysis may be inducing apoptosis. Following specific drug combination treatment, 2 x

106 cells will be washed 2x with PBS. 1mL diluted FITC-Annexin V (1 µg/mL) in

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binding buffer will be used to resuspend cells. This will incubate for 10 min at RT in the dark.10 µL propidium iodide solution will be added to a final concentration of 1 µg/ml prior to analysis via flow cytometry.

6.5.3 Anticipated results

Akin to sorafenib, we anticipate that other therapies targeting

VEGF/tyrosine kinase signaling (axitinib, sunitinib, pazopanib and bevacizumab) may show significantly higher anti-proliferative and anti-metastatic properties when treated with PTX in RCC cell models, particularly ACHN. In addition, the mTOR inhibitors everolimus and temsirolimus cause p4EBP1 decrease and may yield significant effect on p4EBP1 loss and cyclin D1 with PTX treatment.

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APPENDIX A: DIFFERENTIAL MODULATION OF p-4EBP1, p-ERK AND CYCLIN D1 BY AMUVATINIB/ERLOTINIB TREATMENT IN PTEN+ AND PTEN- PROSTATE CANCER CELLS

A.1 Abstract

PTEN negatively regulates the PI3K/AKT/mTOR pathway, and the PTEN gene is mutated/deleted in ~60% of prostate cancer cases. Downstream of mTOR, eukaryotic initiation factor 4E binding protein 1 (4EBP1) regulates cyclin

D1 translationally. In contrast, ERK, a member of the MAPK pathway, regulates cyclin D1 transcriptionally. Both the PI3K and MAPK signaling pathways contribute to disease progression in prostate cancer. A PTEN-deficient human prostate cancer cell line, LNCaP, and a PTEN-positive human prostate cancer cell line, DU145, were used to examine the role of PTEN status in determining drug sensitivity to amuvatinib, a receptor tyrosine kinase inhibitor, and erlotinib, an epidermal growth factor inhibitor. Combination drug treatment in LNCaP caused a decrease in p-4EBP1 (Ser65, Thr70 and Thr37/46), with a concomitant decrease in cyclin D1 protein levels. 2D Western and MALDI-TOF mass spectrometry also revealed loss of hyperphosphorylation of 4EBP1 following combination treatment. Moreover, amuvatinib alone, but not erlotinib, decreased p-4EBP1 and cyclin D1. In DU-145 cells, combination drug treatment had no effect on p-4EBP1 status, but decreased p-ERK and cyclin D1 protein levels, concomitantly. Additionally, erlotinib alone, but not amuvatinib, caused a loss of p-ERK and cyclin D1. The data reveal the differential modulation of cyclin D1 protein levels, and its upstream regulators, in the PTEN- and PTEN+ cell lines by 173

amuvatinib and erlotinib. Such differential responses likely contribute to specific anti-tumor drug efficacy. Understanding cell-specific modulation of cyclin D1 will assist in the selection of appropriate biomarkers to assist in directing patient- targeted therapy.

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A.2 Introduction

Prostate cancer is the second most diagnosed cancer and the second leading cause of cancer death among men in the United States (Siegel et al.,

2014). Activity of the tumor suppressor, phosphatase and tensin homolog

(PTEN), is lost in about 60% of prostate cancer cases (Cairns et al., 1997;

Whang et al., 1998). PTEN negatively regulates the PI3K (phosphoinositide 3- kinase)/AKT/mammalian target of rapamycin (mTOR) pathway, which is critical for prostate cancer progression and correlates with poor clinical outcome (Graff et al., 2000; Kreisberg et al., 2004; Le Page, et al., 2006; Liao et al., 2003; Sarker et al., 2009). In addition, the mitogen activated protein kinase (MAPK) pathway has been implicated in prostate cancer progression (Ahn et al., 2013; Chen et al.,

2013; Gioeli et al., 1999; Li et al., 2014; McCubrey et al., 2007; Park et al., 2014).

Diverse PTEN and MAPK signaling heterogeneity within particular prostate cancer tumors have been reported (Malik et al., 2002; Yoshimoto et al., 2006), demonstrating the importance of both signaling pathways in prostate cancer progression and, thus, therapeutic targeting.

Cyclin D1 is required for progression from the G1 phase into the S phase of the cell cycle (Sherr, 1996). Over-expression of cyclin D1 causes an increase in cell cycle progression and cell proliferation, implicating it in a variety of cancers including prostate cancer(Chen et al., 1998; Drobnjak et al., 2000; Ju et al.,

2014). Downstream of the PI3K/AKT/mTOR pathway, eukaryotic initiation factor

4E (eIF4E) binding protein 1 (4EBP1), regulates cyclin D1 expression through

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binding and inhibiting eIF4E mediated 5' cap-dependent translation (De Benedetti and Graff, 2004). Hyperphosphorylation of 4EBP1 (Ser 65, Thr 70) via upstream signaling activation causes decreased affinity between 4EBP1 and eIF4E, allowing for translation of cyclin D1 to occur. In contrast, extracellular-signal- regulated kinase 1/2 (ERK), a member of the MAPK pathway, regulates cyclin D1 expression through activation of Activator Protein-1 (AP-1) transcription factor

(Gao et al., 2008).

Amuvatinib (MP470) is a receptor tyrosine kinase (RTK) inhibitor that selectively inhibits the c-Kit/PDGFR/c-Met/AXL RTKs (Qi et al., 2009). It has been shown to be cytotoxic in gastrointestinal stromal tumor cells (Mahadevan et al., 2007), radiosensitizes glioblastoma cells (Welsh et al., 2009) as well as

H1299 lung carcinoma cells (Zhao et al., 2011) and reversed epithelial-to- mesenchymal transition in murine breast cancer stem cells (Asiedu et al., 2014).

Erlotinib (Tarceva) is an epidermal growth factor receptor (EGFR) inhibitor that has gained FDA approval for the treatment of non-small cell lung cancer and pancreatic cancer. Activation of compensatory pathways through cross talk between the MAPK and the PI3K/AKT/mTOR pathways in cancer when treated with single agent therapies have been reported (Aksamitiene et al., 2012; De

Luca et al., 2012). Further, PTEN loss and AKT activation have been reported to contribute to erlotinib resistance in lung cancer cells (Sos et al., 2009). These studies stress importance of dual inhibition of MAPK and PI3K pathways in treating prostate cancer. Combined inhibition of these pathways blocks cell proliferation and leads to BCl-2 and Bim upregulation in pre-clinical samples 176

(Hour et al., 2012; Kinkade et al., 2008). Additionally, PI3K and MAPK treatment in patients with advanced cancer suggested dual inhibition may potentially exhibit favorable efficacy compared to single agent intervention (Shimizu et al., 2012)

Previously, we reported that amuvatinib, in combination with erlotinib, demonstrated enhanced cytotoxicity and induction of apoptosis in prostate cancer cells as well as tumor growth inhibition of LNCaP xenografts in SCID mice

(Qi et al., 2009).

In this study the PTEN-deficient human prostate cancer cell line, LNCaP, and the PTEN-positive human prostate cancer cell line, DU145, were used to examine the role of PTEN-status in determining drug sensitivity to amuvatinib and/or erlotinib with regard to decreased cyclin D1 protein levels as well as modulation of the upstream MAPK and PI3K pathways. We examined the effect of amuvatinib and erlotinib, dosed separately and in combination, to understand molecular mechanisms that may help to describe the benefit of combination dose reported previously (Qi et al., 2009). Improved understanding of the molecular events underlying prostate carcinogenesis, and how they affect specific drug sensitivity, may provide further insight into personalizing targeted cancer therapies in patients.

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A.3 Materials and Methods

A.3.1 Cell Culture

LNCaP and DU-145 cells (ATCC, Manassas, Virginia) were grown in

RPMI 1640 and DMEM (Corning, Manassas, Virginia), respectively, containing

˚ 10% FBS. Cells were grown at 37 C in a humidified atmosphere of 5% CO2.

A.3.2 Immunohistochemistry

Paraffin-embedded tissue array blocks were sectioned at 4 µm. Slides were heated for 1 hr at 60ºC deparaffinized in xylene, rehydrated in graded alcohol, and rinsed in distilled water. Antigen retrieval and immunohistochemistry were performed using either a Benchmark XT or Discovery XT automated immunohistochemistry system (Ventana Medical Systems, Inc., Tucson, AZ) using conditions which were validated for the 4EBP1 antibody. Tissue array slides were stained with primary antibodies against 4EBP1 (Novus Biologicals,

Littleton, CO). Primary antibodies were detected using iView DAB orDABMAP

(Ventana Medical Systems, Inc.) and tissue was counterstained in hematoxylin.

A.3.3 1D/2D Western Blot Analysis

LNCaP and DU-145 cell lysates were generated using Cell Lysis Buffer

10X (Cell Signaling Technologies, Danvers, MA) containing 1 mM Pefabloc SC,

Complete protease inhibitor cocktail tablet and phosphatase inhibitor cocktail

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tablet (Roche, South San Francisco, CA). Protein concentration was determined with the DC Protein Assay (Bio-Rad, Hercules, CA). 1D Western: 30-50µg protein was subjected to 10, 12 or 14% SDS-PAGE at 140V, followed by electrophoretic transfer to PVDF membranes. 2D Western: 10µg protein was loaded onto pH 4-7 ReadyStrip IPG IEF strip (Bio-Rad), focused for 1 hour at

500V, and then run at 8,000V for a total of 45,000 V/hrs. Focused strips were then incubated with 4 mL DTT/ equilibrium buffer (150mM tris-HCl, 6M urea, 30% glycerol, 2% SDS, 10 mg/mL DTT) for 15 minutes, washed twice with distilled water and incubated in the dark with 4mL iodoacetamide/equilibrium buffer

(150mM tris-HCl, 6M urea, 30% glycerol, 2% SDS,25 mg/mL iodoacetamide) for

15 minutes. The strips were then run on a Criterion TGX IPG 4-20% SDS-PAGE gel (Bio-Rad) for 55min at 200V, and electrophoretically transferred to PVDF membranes. Primary antibodies used were p-ERK (4376), ERK, p-AKT, AKT, cyclin D1 (2926), p-4EBP1 Ser65 (9451), Thr70 (5078) and Thr37/46 (2855) and

4EBP1 (9452) (Cell Signaling Technologies), and GAPDH (8245) (Abcam,

Cambridge, MA). Secondary anti-mouse and anti-rabbit immunoglobulin conjugated with horseradish peroxidase (Santa Cruz Biotechnology, Dallas,

Texas) were used at a 1:3000 dilution in 5% milk/TBS-T. The blots were visualized with ECL Western Blotting Detection Reagent (Thermo, Waltham,

MA).

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A.3.4 4EBP1 Immunoprecipitation

LNCaP total cell lysate (1 mg) tumbled with 10 µg mouse 4EBP1 monoclonal antibody (Cell Signaling Technologies) for 1 hr at 4 ºC. 25 µL protein

G sepharose bead slurry (Amersham, Pittsburgh, PA) was added and incubated with rotation overnight at 4 ºC. The beads were washed thrice with phosphate- buffered saline (PBS). The immunoprecipitated (IP) proteins were eluted from beads with using buffer containing 50 mM DTT and boiled at 95 ºC for 5 min. The

IP was then combined with ice-cold methanol in 1.5 mL Eppendorf tubes and was chilled at -80 C for 1 hr to induce protein precipitation. The IP sample was centrifuged at 13,000g for 5 mins at 4 ºC and supernatant was removed. The IP- methanol precipitate pellet was mixed with 4x XT sample loading buffer (BioRad) with 5% β-mercaptoethanol and boiled for 5 min in preparation for 1D Western analysis.

A.3.5 Statistics

Data are expressed as mean averages ± standard error. Mean values were compared using a Student's T-test, two tails, with equal variances. P-values are expressed as *p<0.05, **p<0.01, and ***p<0.001.

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A.4 Results

A.4.1 LNCaP (PTEN -) display activated PI3K/AKT/mTOR signaling, whereas DU145 (PTEN +) display activated MAPK signaling

The PTEN-deficient cell line, LNCaP, reveals a constitutively activated

PI3K pathway resulting from PTEN activity loss. p-ERK is undetectable in

LNCaP, whereas pAKT and p4EBP1 (Ser 65, Thr 70, and Thr 37/46) levels are high, concomitant with cyclin D1 levels (Figure A.1). The PTEN + cell line,

DU145, primarily express MAPK pathway activation. p-AKT is undetectable, whereas p-ERK levels are high, concomitant with cyclin D1 levels (Figure A.1).

A.4.2 4EBP1 expression is increased in prostate cancer compared to prostatic intraepithelial neoplasia (PIN)

Immunohistochemistry of PIN (Figure A.2A) showed diffuse upregulation of 4EBP1 expression compared to 3+ positive invasive (Gleason grade 3) carcinoma tissue which revealed significantly upregulated levels of 4EBP1

(Figure A.2B).

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Figure A.1 PI3K/AKT/mTOR/4EBP1 and MAPK signaling differences in LNCaP and DU-145 Untreated whole cell lysates were collected from LNCaP and DU-145 cells. Equal amounts of protein were separated via SDS-PAGE and immunoblotted for the targets shown above.

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Figure A.2 4EBP1 levels in PIN and prostate cancer tissues PIN (A) and invasive prostate cancer tissue (Gleason grade 3) with diffuse 3+ cytoplasmic staining (B). 4EBP1 staining is increased in cancer compared to PIN.

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A.4.3 Amuvatinib and combination amuvatinib + erlotinib treatment decreases p4EBP1 and cyclin D1 levels in LNCaP

Combination amuvatinib + erlotinib treatment (10 µM each) (Figure A.3A) decreased p-4EBP1 Ser65 (61, ±6%, ***p<0.001), Thr70 (83, ±1%, ***p<0.001),

Thr37/46 (73, ±13%, **p<0.01) concomitant with cyclin D1 protein level decrease

(52, ±4%, ***p<0.001) in LNCaP at 2.5 hr compared to control. p-ERK levels were undetectable in LNCaP vehicle (Figure A.1) as well as treated samples

(data not shown). Amuvatinib (10 µM) and combination amuvatinib + erlotinib treatment (10 µM each) (Figure A.3B) decreased p-4EBP1 Ser65, Thr70,

Thr37/47, concomitant with cyclin D1 decrease in LNCaP at 2.5 hr compared to control. Control samples received DMSO for 2.5 hr.

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Figure A.3 The effects of single and combination erlotinib (10 μM) and amuvatinib (10 μM) treatment in LNCaP

(A) Western blot analysis of indicated proteins following a time-course (0 - 2.5 hr) treatment of combination erlotinib + amuvatinib (10μM each) in LNCaP. (B) Western blot analysis of indicated proteins following treatment with erlotinib (10 μM), amuvatinib (10 μM), or erlotinib + amuvatinib (10 μM each) at 2.5 hr in LNCaP. Control samples received DMSO for 2.5 hr. Data are from a single experiment, representative of three separate experiments.

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A.4.4 Erlotinib and combination erlotinib + amuvatinib treatment decreases pERK and cyclin D1 levels in DU145

Combination erlotinib + amuvatinib treatment (10 µM each) (Figure A.4A) decreased p-ERK (89, ±3%, ***p<0.001) concomitant with cyclin D1 protein decrease (72, ±2%, ***p<0.001) in DU-145 at 2.5 hr compared to control. No modulation was seen in p-4EBP1 levels. Erlotinib (10 µM) and combination erlotinib + amuvatinib treatment (10 µM each) (Figure A.4B) decreased p-ERK

(87%, ±1%, ***p<0.001 and 85%, ±1%, ***p<0.001, respectively), concomitant with cyclin D1 decrease in DU-145 at 2.5 hr compared to control. Control samples received DMSO for 2.5 hr.

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Figure A.4 The effects of single and combination erlotinib (10 μM) and amuvatinib (10 μM) treatment in DU145

(A) Western blot analysis of indicated proteins following a time-course (0 - 2.5 hr) treatment of combination erlotinib + amuvatinib (10μM each) in DU145. (B) Western blot analysis of indicated proteins following treatment with erlotinib (10 μM), amuvatinib (10 μM), or erlotinib + amuvatinib (10 μM each) at 2.5 hr in DU145. Control samples received DMSO for 2.5 hr. Data are from a single experiment, representative of three separate experiments.

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A.4.5 Erlotinib and amuvatinib cause multiple phosphorylation loss of 4EBP1

Whole cell lysate was run on two-dimensional gel electrophoresis and immunoblotted for total 4EBP1 (Figure A.5A). The DMSO sample (Figure A.5A upper) reveals several punctate spots near the lower-pH region (pI: 4.0 - 5.0), signifying higher phosphorylation status. There is also a slight shift towards a higher molecular weight in this region, further signifying hyperphosphorylation.

Combination amuvatinib + erlotinib treatment (10 µM each) (Figure A.5A lower) reveals spots corresponding to phosphorylation loss (pI: 5 - 5.5). Figure A.5B shows theoretical pI values corresponding to consecutive addition of phosphate groups to 4EBP1. Spots in the DMSO sample (Figure A.5A upper) reflect pI's corresponding to high phosphorylation status (pIs: 4.03, 4.15, 4.27, 4.40, 4.53,

4.67). Spots in the combination drug treated amuvatinib + erlotinib (10 µM each) sample (Figure A.5A lower) reflect pIs corresponding to low (pIs: 4.84, 5.04) or no (pI: 5.32) phosphorylation. One-dimensional Western blot analysis of LNCaP whole cell lysate as well as 4EBP1 IP sample (Figure A.5C) confirmed higher molecular weight 4EBP1 is present in the DMSO whole cell lysate and IP LNCaP samples. Upon combination amuvatinib + erlotinib treatment, 4EBP1 molecular weight decreases in both whole cell lysate as well as the IP sample.

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Figure A.5 Combination erlotinib (10 μM) and amuvatinib (10 μM) treatment decreased 4EBP1 phosphorylation in LNCaP at 2.5 hr (A) Whole cell lysate (wcl) from LNCaP samples treated with DMSO or combination amuvatinib (10 µM) + erlotinib (10 µM) for 2.5 hr was loaded onto a pH 4-7 IEF strip and focused, then run on two-dimensional gel electrophoresis and immunoblotted for total 4EBP1 protein. (B) Theoretical pI values corresponding to addition of phosphate groups to 4EBP1. (C) One-dimensional Western blot analysis of LNCaP whole cell lysate as well as 4EBP1 IP sample following combination treatment for 2.5 hr immunoblotted for total 4EBP1 protein. Figures are from the same blot with lanes removed for clarity. Data are from a single experiment, representative of three separate experiments.

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A.5 Discussion

Prostate cancer signaling heterogeneity is an important concept in determining optimal treatment strategies. PI3K pathway activation, presumably through PTEN loss, has been significantly linked to prostate cancer progression and correlates with poor clinical outcome. In addition, the MAPK signaling pathway has been implicated in the progression of prostate cancer. MAPK and

PTEN pathway activation are not always mutually exclusive, as it was shown in

35 prostate cancer tumors via FISH and IHC analysis that PTEN was lost in a range of 0 - 97% of cells within each tumor sample. There is dispute, however, regarding how the respective MAPK and PI3K signaling pathways affect prostate tumor progression. Tumor progression was shown to be accompanied by increased expression of p-AKT, which coincided with suppression of apoptosis

(Paweletz et al., 2001). Further, these studies showed p-ERK was suppressed with progression of disease. These findings are in contrast to studies reporting that increased expression of p-MAPK was associated with increased Gleason score as well as found in recurrent tumors following androgen ablation therapy.

Both MAPK and PI3K pathways appear important in prostate cancer progression and, thus, dual therapeutic targeting of both pathways in the treatment of prostate cancer should be considered.

Using LNCaP (PTEN -) and DU145 (PTEN +) (Figure A.1) we determined how PTEN status affected sensitivity to amuvatinib and erlotinib, as single agents as well as combined (see Figure A.6 for summary). LNCaP responded positively

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to amuvatinib as well as combination amuvatinib + erlotinib treatment via a decrease in p-4EBP1 and cyclin D1 levels (Figure A.3A, B). DU145 responded positively to erlotinib as well as erlotinib + amuvatinib treatment as demonstrated by decrease in p-ERK and cyclin D1 levels (Figure 4A, B). A goal of ours was to understand the molecular mechanism of action exerted by erlotinib and amuvatinib since the combination of the two drugs previously produced remarkable synergistic effects in controlling LNCaP tumor growth when compared to single drug usage.

LNCaP expresses constitutively hyperphosphorylated 4EBP1 at several putative sites, rendering 4EBP1 unable to bind and inhibit eIF4E. This ensures the ability of eIF4E to initiate the translation of proteins believed to contribute to tumor growth and progression (e.g. cyclin D1). The precise pattern of phosphorylation(s) that govern the activity of 4EBP1, however, is unclear. In order to visualize potential drug modulation of both known and possibly unknown phosphorylation sites on 4EBP1, we analyzed combination treated LNCaP whole cell lysate via 2D Western blot analysis (Figure A.5A). Following treatment, immunopositive spots for 4EBP1 displayed a shift in pI, reflective of an approximate loss of 7-8 phosphorylations (Figure A.5B). This suggests combination treatment induces loss of phosphorylation at residues in addition to those specifically analyzed via Western blot (Ser 65, Thr 70 and Thr 37/46;

Figures A.4, A.5). Further, whole cell lysate and IP of 4EBP1 from combination treated LNCaP demonstrated a decrease in molecular weight via 1D western blot analysis, as compared to vehicle control (Figure A.5C). This also demonstrates 191

significant loss of post-translational modification resulting in decreased molecular weight.

Understanding the effects of treatment with erlotinib and amuvatinib, with regard to cyclin D1 decrease, serve as a proof of concept in demonstrating the increased efficacy in using combination therapies. Combining therapeutics can help in surmounting resistance to single agents via compensatory mechanisms, as is often found in recurrent and advanced tumors. Further, combinational treatment can aid in decreasing the toxicity of one or all agents being used. In addition, combination drug therpay can yield synergistic effects, as demonstrated in these findings. Future studies in our laboratory will include investigation into single vs combination erlotinib and amuvantinib treatment in LNCaP and DU-145 xenograft models, in an attempt to replicate our findings in vivo.

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Figure A.6 Proposed signaling and drug perturbation schematic in DU- 145 (PTEN +) and LNCaP (PTEN -) Single treatment with erlotinib caused cyclin D1 decrease in DU-145, whereas single treatment with amuvatinib caused decrease in cyclin D1 in LNCaP. Combination treatment also caused decreased cyclin D1 levels in these cells. Differential single drug effects within DU-145 and LNCaP cell lines are, presumably, due to differences in the pathways regulating cyclin D1 synthesis (MAPK and PTEN/PI3K/AKT/mTOR).

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