The Role of the Rho GEF Arhgef2 in RAS Tumorigenesis

Home , ARHGEF1, ARHGEF12, ARHGEF2, KRAS, RhoG, RIN1

Jane Cullis

A thesis submitted in conformity with the requirements

for the degree of Doctor of Philosophy

Graduate Department of Medical Biophysics

University of Toronto

Jane Cullis Degree of Doctor of Philosophy, 2013 Graduate Department of Medical Biophysics, University of Toronto

Abstract

Tumorigenesis is driven by the sequential accumulation of genetic lesions within a cell, each which confer the cell with traits that enable its abnormal growth. The result is a mass of dysregulated cells, or tumor, which, upon further mutation, may spread, or metastasize, to other organs of the body. The dissemination of tumor cells makes treatment difficult, and thus confers cancer with its associated lethality. Over the past 30 years, the RAS genes have been critical in teaching us the mechanisms underlying the molecular progression of cancer. RAS is mutated in

33% of all cancers and is often an early event in its stepwise progression1. As a result, the RAS genes are widely accepted as ‘drivers’ or ‘initiators’ of human tumorigenesis. Unfortunately, efforts directed at targeting RAS in the clinic have as of yet been unsuccessful. This has triggered a need to identify genes that are required for RAS tumorigenesis that are therapeutically tractable.

My research has focused on deciphering the potential role of the Rho GEF Arhgef2 in RAS- mediated tumorigenesis. I have found that Arhgef2 is a bona fide transcriptional target of RAS and is upregulated in human tumors harboring RAS mutations. Importantly, depletion of Arhgef2 in RAS-mutated cells inhibits their survival, proliferation, and tumor growth in murine models.

In search of the mechanism underlying the requirement of Arhgef2 in RAS tumorigenesis, I have uncovered a novel function for Arhgef2 as a positive regulator of a central RAS pathway, the

ii mitogen-activated protein kinase (MAPK) pathway. Thus, Arhgef2 is part of a positive feedback loop in which RAS-dependent increases in Arhgef2 expression results in the amplification of

RAS signaling. Moreover, Arhgef2 confers tumor cells with properties favoring their malignant conversion, thereby implicating Arhgef2 in the formation of metastases. Together, these studies suggest that Arhgef2 plays an important role at multiple stages of tumorigenic progression and may therefore be a promising therapeutic target in RAS-mutated tumors.

1Karnoub et al., 2008

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Acknowledgments

First and foremost, I would like to thank my amazing family for their unwavering support and encouragement throughout my degree. My father, who not only inspires me with his science, but whose compassion and sense of fun I admire and strive to emulate. My mother, one of the strongest, most brilliant women I know: thank you for being not just a mother to me in the last 6 years but also one of my best friends. My brother Jepray, you have been a tremendous source of comfort, positivity and wisdom and I thank you for always being there for me.

To my supervisor, Rob. You made it about more than science. You are the one who taught me how to run marathons. You taught me that the work you put in is the work you get out; that strength and endurance takes time and patience and cannot be forced; that the lows are worth the highs; that there are no shortcuts; that it’s not how fast you can sprint but how well you can push to the very end. Most importantly, you taught me never to give up until you’re there. Not many people can teach such hard lessons while expressing so much love, compassion and understanding, but you did. Thank you.

Thank you to my committee members, Jane and Dr. Medin. I appreciate the time you took to oversee my project and improve my research with your excellent advice and encouragement.

Thank you to my dear collaborators, Nikolina Radulovich and Dr. Ming Tsao, for your help and guidance with my animal and immunohistochemical studies.

Dedi. You gave meaning to the word ‘Dedidit’ and together, we did it . You are a remarkable scientist but an even more amazing person and friend. Thank you for always putting things in perspective and for making science fun.

Mauricio and Tim, my BFFs OMG. Mauricio, it is largely because of you that I was able to run Rob’s marathons. Thank you for being a constant source of support and fun during the last six years. You have made the difficult times bearable and the good times unbelievable.

To all the members of the Rottapel lab for putting up with me and all my Western blots. Thank you for your encouragement and sense of humor – it is you guys that made coming to lab every day worth it, successful experiment or not. Also, thank you to the ladies from the Kislinger lab,

iv especially Lusia, for either keeping me sane in the office or making the choice to go insane with me.

To my training partners and running friends, the Angels. Thank you for constantly reminding me that there’s more to life than the lab (and for inspiring me to run real marathons). Nic, DocZ, MamaK and Jebs, thank you for your patience, wisdom and guidance in all aspects of life.

To the others along the way that have inspired and encouraged me – Delilah (a.k.a. Topicoolis) and my beautiful cousin Sarika – you are two of the most important people in my life and it’s been so comforting knowing you were always there for me. You have each helped me in such different but crucial ways and I can’t thank you enough.

Finally, I absolutely have to thank Goose for keeping me going during the last six years. You taught me to relax, gave me the energy to keep going and were always there when I needed you. Cheers.

Table of Contents

Abstract ...... ii

Acknowledgments...... iv

Table of Contents ...... vi

List of Tables ...... ix

List of Figures ...... x

List of Appendices ...... xii

Appendix 1: Microarray Analysis of PANC-1 and H-RASV12-Transformed Fibroblast Cells Harboring Stable Arhgef2 Knockdown...... xii

List of Abbreviations and Symbols...... xiv

Chapter 1 ...... 1

Introduction ...... 1

1.1 The RAS superfamily of small GTPases ...... 1

1.2 The RAS subfamily ...... 4

1.3 The Rho subfamily ...... 9

1.4 Guanine nucleotide exchange factors ...... 14

1.5 Arhgef2...... 19

Chapter 2 ...... 25

Arhgef2 Provides a Positive Feedback Loop Required for Signaling Through the Oncogenic RAS Pathway ...... 25

2.1 Abstract ...... 25

2.2 Introduction ...... 26

2.3 Experimental Procedures...... 29

2.4 Results ...... 36

2.4.1 Arhgef2 protein expression is acutely induced by the RAS/MAPK pathway ...... 36

2.4.2 ARHGEF2 is a transcriptional target of the RAS/MAPK pathway ...... 38

2.4.3 Arhgef2 is required for cell survival downstream of oncogenic RAS ...... 40

2.4.4 Arhgef2 contributes to RASV12-mediated cellular transformation in vitro and in vivo 43

2.4.5 Arhgef2 contributes to the increased proliferative capacity of RASV12-transformed fibroblasts in a GEF-independent manner ...... 45

2.4.6 Arhgef2 is required for MAPK pathway activation in response to oncogenic RAS .... 46

2.4.7 Arhgef2 is a component of the KSR-1 complex and is required for the dephosphorylation of its negative regulatory site on S392...... 50

2.4.8 Arhgef2 is required for PP2A-mediated dephosphorylation of KSR-1 on S392 ...... 54

2.5 Discussion ...... 57

Chapter 3 ...... 62

Arhgef2 is Required for Primary Tumorigenesis and Promotes Mesenchymal Transition in Pancreatic Ductal Adenocarcinoma ...... 62

3.1 Abstract ...... 62

3.2 Introduction ...... 63

3.3 Experimental Procedures...... 67

3.4 Results ...... 73

3.4.1 ARHGEF2 is essential across several human epithelial cancer cell lines and its protein expression is regulated by the RAS/MAPK pathway ...... 73

3.4.2 Arhgef2 is required for PDAC tumor growth in vivo ...... 74

3.4.3 Arhgef2 expression correlates with advanced tumor grade in human lung, colorectal and pancreatic cancer ...... 78

3.4.4 Arhgef2 expression alters gene signatures associated with epithelial differentiation state ...... 79

3.4.5 Arhgef2 suppresses the epithelial cell phenotype in RAS-independent human adenocarcinoma cell lines ...... 82

3.4.6 Arhgef2 is required for TGF-induced epithelial-to-mesenchymal-transition in a mammary epithelial cell model ...... 86

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3.5 Discussion ...... 93

Chapter 4 ...... 99

Future Perspectives ...... 99

4.1 Abstract ...... 99

4.2 Experimental Procedures...... 100

4.3 Future Perspectives ...... 102

4.3.1 The role of Arhgef2 in metastases ...... 102

4.3.2 The cooperation of Arhgef2 with mutant p53 ...... 104

4.3.3 The regulation of Arhgef2 by anti-mitotic chemotherapeutic agents ...... 110

4.3.4 Arhgef2 as a therapeutic target ...... 112

Concluding Remarks ...... 116

Appendix ...... 117

Appendix 1: Microarray analysis of PANC-1 and H-RASV12-Transformed Fibroblast Cells Harboring Stable Arhgef2 Knockdown...... 117

Table I Functional annotation clustering of upregulated genes in Arhgef2-depleted PANC-1 cells...... 117

Table II Upregulated gene lists in Arhgef2-depleted PANC-1 cells by functional annotation118

Table III Functional annotation clustering of downregulated genes in Arhgef2-depleted PANC-1 cells...... 123

Table IV Downregulated gene lists in Arhgef2-depleted PANC-1 cells by functional annotation ...... 125

Table V Functional annotation clustering of Significance Analysis of Microarray (SAM) genes downregulated in Arhgef2-depleted NIH 3T3-H-RasV12 cells ...... 128

Table VI Downregulated gene lists in Arhgef2-depleted NIH 3T3-H-RASV12 cells by functional annotation ...... 131

References ...... 133

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List of Tables

Table 2.1 Murine and Human Arhgef2 shRNA and GFP shRNA Sequences

Table 3.1 Gene Target Primer Sequences

Table 4.1 RAS/MAPK and p53 mutations in ARHGEF2-essential cell lines

List of Figures

Figure 1.1 The RAS superfamily of small GTPases Figure 1.2 Small GTPase domain organization Figure 1.3 The GTPase cycle Figure 1.4 RAS isoform mutations in human cancer Figure 1.5 RAS effector pathways Figure 1.6 Crystal structure of the DH-PH domain of Dbl’s big sister (Dbs) in complex with RhoA Figure 1.7 The diversity in Rho GEF domain organization Figure 1.8 The domain organization of Arhgef2 Figure 2.1 Arhgef2 protein expression is acutely induced by oncogenic RAS

Figure 2.1 Arhgef2 protein expression is acutely induced by oncogenic RAS

Figure 2.2 H-RASV12-induced Arhgef2 upregulation is dependent on MAPK pathway activation

Figure 2.3 ARHGEF2 is a transcriptional target of the RAS/MAPK pathway

Figure 2.4 Arhgef2 is required for cell survival downstream of oncogenic RAS

Figure 2.5 Arhgef2 contributes to RASV12-mediated cellular transformation in vitro and in vivo

Figure 2.6 Arhgef2 protein expression is regained in a subset of Arhgef2-knockdown xenografts

Figure 2.7 Arhgef2 contributes to the proliferative capacity of RASV12-transformed fibroblasts in a GEF-independent manner

Figure 2.8 Arhgef2 is required for MAPK pathway activation in response to oncogenic RAS

Figure 2.9 Arhgef2 is a component of the KSR-1 complex and is required for the dephosphorylation of the negative regulatory site Ser392 on KSR-1

Figure 2.10 Arhgef2 is required for plasma membrane translocation of KSR-1

Figure 2.11 Arhgef2 is required for PP2A-mediated dephosphorylation of KSR-1 on S392

Figure 2.12 The Arhgef2/PP2A complex provides a positive feedback loop to the KSR/MAPK pathway in RASV12-transformed cells

Figure 3.1 Multistep tumorigenesis in pancreatic ductal adenocarcinoma

Figure 3.2 ARHGEF2 is essential across several human epithelial cancer cell lines

Figure 3.3 ARHGEF2 protein expression is regulated by the RAS/MAPK pathway in human epithelial cell lines

Figure 3.4 Arhgef2 is required for pancreatic tumor growth in vivo

Figure 3.5 Arhgef2 is required for KSR-1 S392 dephosphorylation, ERK1/2 phosphorylation and proliferation in PDAC cells

Figure 3.6 Arhgef2 expression correlates with advanced tumor grade in human lung, colon and pancreatic tissue microarrays

Figure 3.7 Arhgef2 suppresses the epithelial cell phenotype in human adenocarcinoma cell lines

Figure 3.8 TGF induces epithelial-to-mesenchymal-transition in a normal mammary gland epithelial cell model

Figure 3.9 Arhgef2 is required for TGF-induced epithelial-to-mesenchymal-transition in NMuMG cells

Figure 3.10 Arhgef2 may promote EMT via its cooperative regulation by RASV12, p53 and TGF signaling pathways

Figure 4.1 Arhgef2 is highly expressed in serous ovarian carcinoma Figure 4.2 Arhgef2 is essential for survival in OVCAR5 cells Figure 4.3 Arhgef2 gene expression correlates with essentiality in serous ovarian carcinoma Figure 4.4 Arhgef2 expression in ovarian carcinoma cell lines

List of Appendices

Appendix 1: Microarray Analysis of PANC-1 and H-RASV12-Transformed Fibroblast Cells Harboring Stable Arhgef2 Knockdown Table I Functional annotation clustering of upregulated genes in Arhgef2-depleted PANC-1 cells I.A Biological Process I.B Molecular Function I.C KEGG Pathway Table II Upregulated gene lists in Arhgef2-depleted PANC-1 cells by functional annotation II.A Regulation of Apoptosis II.B Biological Adhesion II.C Cell Motion II.D Cell Junction II.E Focal Adhesion II.F ECM-Receptor Interaction Table III Functional annotation clustering of downregulated genes in Arhgef2-depleted PANC-1 cells III.A Biological Process III.B Cellular Component III.C Molecular Function III.D KEGG Pathway Table IV Downregulated gene lists in Arhgef2-depleted PANC-1 cells by functional annotation IV.A Mesenchymal Cell Development/Differentiation IV.B Anti-Apoptosis IV.C Cell Migration IV.D Cytoskeleton IV.E Focal Adhesion Table V Functional annotation clustering of Significance Analysis of Microarray (SAM) genes downregulated in Arhgef2-depleted NIH 3T3-H-RASV12 cells V.A Biological Process V.B Cellular Component

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V.C Molecular Function V.D KEGG Pathway Table VI Downregulated gene lists in Arhgef2-depleted NIH 3T3-H-RASV12 cells by functional annotation VI.A Response to Wounding VI.B Epithelial Cell Differentiation VI.C Fibroblast Growth Factor Receptor Signaling Pathway VI.D Cell-Matrix Adhesion VI.E Adherens Junctions VI.F Cell Junction VI.G Tight Junction

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List of Abbreviations and Symbols Amino Acids

Ala/A/Alanine; Arg/R/Arginine; Asn/N/Asparagine; Asp/D/Aspartic Acid; Cys/C/Cysteine; Glu/E/Glutamic Acid; Gln/Q/Glutamine; Gly/G/Glycine; His/H/Histidine; Ile/I/Isoleucine; Leu/L/Leucine; Lys/K/Lysine; Met/M/Methionine; Phe/F/Phenylalanine; Pro/P/Proline; Ser/S/Serine; Thr/T/Threonine; Trp/W/Tryptophan; Tyr/Y/Tyrosine; Val/V/Valine

Symbols

 Alpha  Beta  Gamma  Epsilon  Zeta  Eta  3.14 (pi) m milli, 1x10-3 -6  micro, 1x10 -9 n nano, 1x10 °C degrees Celcius g gram L litre M Molar U Units

GENE names denoted by upper case lettering

Abbreviations

ADC Adenocarcinoma ADH Atypical Ductal Hyperplasia AF-6 ALL Fusion partner 6 AKAP A-Kinase Anchoring Protein Akt Ak thymoma ALL Acute Lymphoblastic Leukemia ALN-VSP ALNylam Vascular endothelial growth factor kinesin Spindle Protein AMCD Anti-Mitotic Chemotherapeutic Drug AML Acute Myeloid Leukemia APC Adenomatous Polyposis Coli AR Androgen Receptor ASEF APC-Stimulated guanine nucleotide Exchange Factor ATCC American Type Culture Collection ATF2 Activating Transcription Factor 2 ATP Adenosine TriPhosphate xiv

AurkA Aurora kinase A AurkB Aurora kinase B B Protein phosphatase 2A, 55kDa regulatory subunit B’ Protein phosphatase 2A, 56kDa regulatory subunit, alpha isoform B56 Protein phosphatase 2A, 56kDa regulatory subunit, alpha isoform BAC Bacterial Artificial Chromosome Bcl B cell lymphoma Bcl-2 B cell lymphoma-2 Bcl-XL B cell lymphoma eXtra Large BCR Breakpoint Cluster Region BFA BreFeldin A BLAST Basic Local Alignment Search Tool BLAT BLAST-Like Alignment Tool BrdU BromodeoxyUridine BSA Bovine Serum Albumin C-terminal Carboxy-terminal C1 Zinc finger domain or cysteine rich domain C3 Clostridium Botulinum 3 C57BL/6 C57 BLack 6 CalPhos Calcium Phosphate CAAX Cysteine-Alanine-Alanine-X amino acid CC Coiled Coil Cdc Cell division cycle protein Cdc42 Cell division control protein 42 Cdk Cyclin-dependent kinase cDNA complementary DNA C. elegans Caenorhabditis elegans CH Calponin Homology CIP Calf Intestinal Phosphatase CML Chronic Myelogenous Leukemia CMML Chronic MyeloMonocytic Leukemia CMV CytoMegaloVirus CMV-Cre CytoMegaloVirus Type I topoisomerase CO2 Carbon dioxide (O2) COMe CarbOxyMethylation CR Cysteine Rich cRNA complementary RiboNucleic Acid CST Cell Signaling Technology CT Cycle Threshold D2O Deuterium Oxide (heavy water) DAG DiAcylGlycerol DAVID Database for Annotation, Visualization and Integrated Discovery DBL Diffuse B-cell Lymphoma Dbs Diffuse B-cell Lymphoma’s big sister DH Dbl Homology DMEM Dulbecco’s Modified Eagles Medium

DMSO DiMethylSulfOxide DNA DeoxyriboNucleic Acid DOCK Dedicator Of CytoKinesis dsDNA double stranded DeoxyriboNucleic Acid DTT DithioThreiTol E14K Adenovirus type 2 Early protein, 14KDa ECAD E-CADherin ECM ExtraCellular Matrix ECT2 Epithelial Cell Transforming sequence 2 EDTA EthyleneDiamineTetraacetic Acid eGFP enhanced Green Fluorescent Protein EGF Epidermal Growth Factor EMT Epithelial-to-Mesenchymal-Transition EPEC EnteroPathogenic Escherichia Coli ER Endoplasmic Reticulum ER Estrogen Receptor ERBB2 v-erb-b2 erythroblastic leukemia viral oncogene homolog 2 ERK Extracellular signal-Regulated Kinase ES Embryonic Stem FBS Fetal Bovine Serum FPKM Fragments Per Kilobase of exon per Million fragments mapped FRET Fluorescence Resonance Energy Transfer FTase Farnesyl Transferase FTI Farnesyl Transferase Inhibitor G418 Geneticin/Neomycin GAP GTPase Activating Protein GARP Gene Activity Rank Profile GDI Guanine nucleotide Dissociation Inhibitor G domain GDP/GTP binding domain GDP Guanosine DiPhosphate GDS Guanine nucleotide Dissociation Stimulator GEF Guanine nucleotide Exchange Factor GEF-H1 Guanine nucleotide Exchange Factor-H1 GOF Gain Of Function GPCR G Protein Coupled Receptor GRB2 Growth factor Receptor-Bound protein 2 GTP Guanosine TriPhosphate GTPase Guanosine TriPhosphatase GGTase GeranylGeranyl Transferase H2O Hydrogen DiOxide (water) H3K4me3 Histone 3 Lysine 4 trimethylation HBSS Hank’s Buffered Saline Solution HCC HepatoCellular Carcinoma HEK Human Embryonic Kidney 293 HEPES 4-(2-HydroxyEthyl)-1-PiperazineEthaneSulfonic acid hPTTG1 human Pituitary Transforming Gene 1

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H-RAS Harvey RAt Sarcoma HRP HorseRadish Peroxidase Hsc70 Heat shock cognate protein 70 Hsp90 Heat shock protein 90 HSQC Heteronuclear Single Quantum Coherence HV HyperVariable region ICMT1 IsoprenylCysteine O-MethylTransferase 1 IF ImmunoFluorescence IgG Immunoglobulin G IHC ImmunoHistoChemistry IKK I appa-B Kinase IP ImmunoPrecipitate kDa kiloDalton i.v. intravenous JNK c-Jun N-terminal Kinase Kb Kilobase KEGG Kyoto Encyclopedia of Genes and Genomes K-RAS Kirsten RAt Sarcoma K-RAS4A Kirsten RAt Sarcoma, isoform 4A K-RAS4B Kirsten RAt Sarcoma, isoform 4B KSP Kinesin Spindle Protein KSR-1 Kinase Suppressor of RAS-1 KO KnockOut LARG Leukemia Associated Rho GEF LBC Lymphoid Blast Crisis LCUC Large Cell Undifferentiated Carcinoma of the lung LFC LBC’s First Cousin LNP Lipid NanoParticle loxP lox sequence derived from bacteriophage P1 LPA Lysophosphatidic Acid LSC LBC’s Second Cousin LY LY294002 MAP Microtubule Associated Protein MAPK Mitogen Activated Protein Kinase MAPKK Mitogen Activated Protein Kinase Kinase MAPKKK Mitogen Activated Protein Kinase Kinase Kinase MDR-1 MultiDrug Resistance-1 MEF Murine Embryonic Fibroblast MEK Mitogen activated protein kinase ERK Kinase Mg+2 Magnesium MgCl2 Magnesium Chloride MLL Mixed Lineage Leukemia mRNA messenger RiboNucleic Acid Myc MyeloCytomatosis mTOR mammalian Target of Rapamycin N-RAS Neuroblastoma RAt Sarcoma

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N-terminal amino-terminal NaCl sodium (Na) Chloride NaF sodium (Na) Fluoride Na3VO4 sodium (Na) orthovanadate (VO4) NET1 NeuroEpithelial cell Transforming 1 NF-1 GAP NeuroFibromatosis-1 GTPase Activating Factor NFB Nuclear Factor kappa () B NIH National Institute of Health NMR Nuclear Magnetic Resonance NMuMG Normal Murine Mammary Gland NSCLC Non Small-Cell Lung Cancer OC Ovarian Carcinoma OCT Optimal Cutting Temperature OHT Hydroxy Tamoxifen OVCAR OVarian CARcinoma p120RASGAPRAS GTPase activating protein, 120kDa p190RhoGAP Ras HOmology GTPase Activating Protein, 190kDa p21 cyclin-dependent kinase inhibitor 1 p27 cyclin-dependent kinase inhibitor 1b PAK P21 Activated Kinase PanIN Pancreatic Intraepithelial Neoplasia PBD P-21 Activated Kinase Binding Domain PBS Phosphate Buffered Saline PBST Phosphate Buffered Saline Tween PCR Polymerase Chain Reaction PD PD98059 PDAC Pancreatic Ductal AdenoCarcinoma PDGF Platelet-Derived Growth Factor PDZ Postsynaptic density protein (PSD95), Drosophila disc large tumor suppressor (Dgl1), Zona Occludens protein-1 (ZO-1) PH Pleckstrin Homology Pi inorganic Phosphate PI3K PhosphoInositide 3-Kinase PIP PhosphatidylInositol Phosphate PKA Protein Kinase A PKB Protein Kinase B PKC Protein Kinase C PLC PhosphoLipase C PMSF PhenylMethaneSulphonylFluoride PP2A Protein Phosphatase 2A PP2Ac Protein Phosphatase 2A catalytic subunit PPP2R2A Protein Phosphatase 2A, 55kDa Regulatory subunit, alpha isoform PPP2R5A Protein Phosphatase 2A, 56kDa Regulatory subunit, alpha isoform PPP2R5B Protein Phosphatase 2A, 56kDa Regulatory subunit, beta isoform PPP2R5E Protein Phosphatase 2A, 56kDa Regulatory subunit, eta isoform PR55 Protein phosphatase 2A, 55kDa regulatory subunit

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PR65 Protein phosphatase 2A, 56kDa regulatory subunit, alpha isoform P-REX Phosphatidylinositol 3,4,5-triphosphate-dependent Rac nucleotide Exchanger pS phosphoSerine PSA Prostate Specific Antigen pT phosphoThreonine PTase PalmitoylTransferase PTEN Phosphatase and TENsin homolog PVDF PolyVinyliDene Fluoride qPCR quantitative Polymerase Chain Reaction R2 Protein phosphatase 2A, 55kDa regulatory subunit R5 Protein phosphatase 2A, 56kDa regulatory subunit, alpha isoform Rac RAS-relAted C3 botulinum toxin substrate RacGEF RAS-relAted C3 botulinum toxin substrate Guanine nucleotide Exchange Factor RAS RAt Sarcoma RASGEF RAt Sarcoma Guanine nucleotide Exchange Factor RASGRF RAS Guanosine Releasing Factor RASSF RAS ASSociation domain Family RBD Rhotekin Binding Domain RCE1 RAS-Converting Enzyme 1 RGS Regulator of G protein Signaling Rho RAS homology RhoA RAS homology, member A RhoB RAS homology, member B RhoC RAS homology, member C RhoE RAS homology, member E RhoGDI RAS homology Guanine nucleotide Dissociation Inhibitor RhoGEF RAS homology Guanine nucleotide Exchange Factor RIN-1 RAS and Rab INteractor – 1 RNA RiboNucleic Acid RNAi RNA interference RNASeq RiboNucleic Acid whole transcriptome shotgun Sequencing ROCK RhO-associated Kinase RPMI Roswell Park Memorial Institute medium RTK Receptor Tyrosine Kinase SAM Significance Analysis of Microarrays SAP-1 Serum Response Factor Accessory Protein-1 SCID Severe Combined ImmunoDeficiency SD Standard Deviation SDS-PAGE Sodium Dodecyl Sulfate-PolyAcrylamide Gel Electrophoresis SE Standard Error Ser/Thr Serine/Threonine SH2 Src Homology 2 SH3 Src Homology 3 shARP small hairpin Activity Ranking Profile shGEF small hairpin against ArhGEF2 shGFP small hairpin against Green Fluorescent Protein

xix shRNA small hairpin RiboNucleic Acid SI Switch I SII Switch II siRNA small interfering RiboNucleic Acid SMD1 Small nuclear ribonucleoprotein 1 SOC Serous Ovarian Carcinoma SOS Son Of Sevenless SQ SQuamous SRC SaRComa SRF Serum Response Factor STRN3 Striatin, calmodulin binding protein 3 (Protein Phosphatase 2A B’’’ subunit) TBK1 Tank-Binding Kinase 1 TBS-T Tris-Buffered Saline Tween TCEP Tris[2-CarboxyEthyl]Phosphine Tctex-1 T-complex testis-specific protein-1 TGF Transforming Growth Factor beta TGH Toronto General Hospital TIAM1 T-cell lymphoma Invasion and Metastasis-Inducing 1 TMA Tumor MicroArray TNF Tumor Necrosis Factor alpha TRC The Ribonucleic acid i Consortium TRIO TRIple Functional dOmain protein TrkBT1 Tyrosine-related kinase B Truncated I Tris Tris(hydroxymethyl)aminomethane TSS Transcription Start Site UO UO126 UTR UnTranslated Region VEGF Vascular Endothelial Growth Factor VIM VIMentin vs versus VSV-g Vesicular Stomatitis Virus-glycoprotein Waf1 Wild-type p53 activated fragment 1 WB Western Blot WCL Whole Cell Lysate WT Wild Type ZEB-1 Zinc finger E-Box-binding homeobox 1 ZO Zona Occludens ZONAB ZONa occludens Associated Y Box factor

Chapter 1 Introduction 1.1 The RAS superfamily of small GTPases

The RAS GTPases are pleiotropic signaling molecules that regulate most core biochemical processes in the cell. They are binary switches that cycle between an inactive, GDP-bound state and an active, GTP-bound state and function as signaling nodes, linking multiple extracellular stimuli to a wide range of intracellular signaling pathways (Vetter and Wittinghofer, 2001). The RAS superfamily comprises 156 members in humans that are divided into five main families based on their sequence and functional similarities: RAS, Rho, Arf, Ran, and Rab (Figure 1.1).

Figure 1.1 The RAS superfamily of small GTPases. The RAS superfamily consists of 156 members in mammals, divided into Arf (27 members), Rab (61 members), Rho (22 members), RAS (36 members) and Ran (1 member) families. The RAS family forms the phylogenetic root of the superfamily and regulates diverse cellular processes including cell proliferation, differentiation, morphology and survival (Karnoub and Weinberg, 2008). The Rho family is predominantly involved in the regulation of the actin cytoskeleton, thereby affecting cell morphology, polarity and migration, and also influences gene transcription and cell cycle progression (Heasman and Ridley, 2008). The Rab and Arf families

2 mainly participate in vesicular cargo trafficking, endocytosis and secretory pathways, however Arf GTPases are distinguished by an additional functional role on microtubules (Zerial and McBride, 2001, Wennerberg et al., 2005). The Ran family is primarily involved in nuclear transport but can also regulate mitotic spindle organization (Clarke et al., 2008).

While functionally diverse, the RAS superfamily members display highly conserved GDP/GTP binding domains, or G domains, which underly their structural similarities and common biochemical properties (Bourne et al., 1991). The G domain is composed of five consensus sequence elements involved in binding phosphate and magnesium or guanine. The GDP/GTP switch mechanism involves a guanine nucleotide-dependent conformational change in two of the G domain sequence elements known as switch I and switch II (Figure 1.2). The interaction with GTP results in a shift in the switch I domain to a position favouring the binding of effector molecules, thereby enabling the activation of downstream signaling pathways (Bishop and Hall, 2000).

Figure 1.2 Small GTPase domain organization. The RAS GTPases exhibit highly conserved G domains that are involved in GTP binding and hydrolysis. The G domain is defined by five consensus sequence elements involved in binding phosphate/Mg2+ (PM) or guanine (G). The switch I (SI) and switch II (SII) regions are critical for GDP/GTP exchange and subsequent effector binding. RAS family members diverge in their C-terminal tails, with RAS and Rho proteins containing hypervariable (HV) regions that dictate their posttranslational modifications. These include prenylation (P), farnestylation (F) and geranylation (G) and subsequent carboxymethylation (C-OMe). The Rab family is commonly modified by geranylation at its C-terminus, while Arf can be myristoylated at its N-terminus to facilitate membrane interactions. Ran is not posttranslationally modified but contains a C-terminal extension required for its activity. The Rho GTPases are distinguished by a 13 amino acid insert within the G domain that is involved in its interaction with effectors (adapted from Vigil et al., 2010). While small GTPases display high affinity for GDP and GTP, they exhibit low intrinsic GTP hydrolysis and inefficient GDP/GTP exchange activities (Bernards and Settleman, 2004). The GDP/GTP switch is determined by two main classes of regulating proteins; guanine nucleotide

3 exchange factors (GEFs), which catalyze the exchange of GDP for GTP, and GTPase activating proteins (GAPs), which promote the intrinsic hydrolysis of GTP (Schmidt and Hall, 2002) (Figure 1.3). The Rho and Rab families are subject to a third class of regulatory proteins known as Rho guanine nucleotide dissociation inhibitors (RhoGDIs), which sequester GDP-bound GTPases in the cytoplasm by masking their lipid membrane-targeting moieties (Olofsson, B, 1999). While the mechanism of action of GEFs, GAPs and GDIs is preserved within each respective class, they display both distinct and shared selectivity for GTPases within each RAS family. This specificity in regulation, combined with the pleiotropy in effectors activated by the GTPases themselves, contributes to their diversity in cellular effects.

Figure 1.3 The GTPase cycle. RAS GTPases cycle between an active, GTP-bound state and an inactive, GDP- bound state. GTPase-activating proteins (GAPs) stimulate their intrinsic hydrolysis of GTP, while guanine nucleotide exchange factors (GEFs) catalyze the exchange of GDP for GTP. Guanine nucleotide dissociation inhibitors (GDIs) act on the Rho and Rab families of RAS GTPases and sequester them in their GDP-bound state. RAS-GTP can bind to its downstream effectors and influence a diverse array of biological processes, including cell proliferation, survival, differentiation, morphogenesis, motility, migration, polarity, gene transcription, vesicle trafficking, nuclear transport, endocytosis and microtubule dynamics.

1.2 The RAS subfamily

The Ras sarcoma (RAS) family of proteins constitute the founding members of the RAS superfamily. They have been the subject of intense study since their discovery in the 1980s due to their high frequency of mutations in human tumors (Bos et al., 1989). The H- and K-RAS genes were first identified in rat cells as retrovirally-transduced oncogenes derived from Harvey and Kirsten sarcoma viruses, respectively, and homologous forms in mouse and human were isolated soon after (DeFeo et al., 1981, Ellis et al., 1982, Chang et al., 1982). Mutant forms of H- and K-RAS were subsequently found in many human cancer cell lines, including those of the bladder, colon and lung (Parada et al., 1982, Der et al., 1982, McBride et al., 1982). Sequencing analysis revealed that the oncogenic forms of H- and K-RAS commonly harbored single point mutations in codon 12, with mutations in codons 13 and 61 less frequently found (Reddy et al., 1982, Taparowsky et al., 1982). A third RAS family member, N-RAS, was isolated in a neuroblastoma cell line and found to contain parallel mutations in codon 12 in human tumors (Hall et al., 1983, Brown et al., 1984).

Localized point mutations in codon 12 of the RAS genes predominantly results in the substitution of a Glycine (G) for a Valine (V) or Aspartic acid (D) and the constitutive activation of the GTPase (Tabin et al., 1982). Research performed in the McCormick laboratory revealed that the active mutants exhibited a three hundred-fold lower GTPase activity compared to their wild-type counterparts, thereby locking them in a GTP-bound state (Clark et al., 1985, Trahey et al., 1987). The oncogenic capacity of RASV12/D12 mutants was demonstrated in focus-forming assays in murine and rodent cells, which underwent morphological transformation in response to their overexpression with cooperating oncogenes (Land et al., 1983, Newbold et al., 1983, Ruley et al., 1983). The physiological significance of RASV12/D12 mutations was appreciated when they were found endogenously in experimental models of carcinogenesis, with H-RASV12 identified in carcinogen-induced mammary and skin tumors, N-RASD12 in thymomas and K-RASD12 in mice exposed to ionizing radiation (Sukumar et al., 1983, Balmain et al., 1983, Guerrero et al., 1984a and b). The most compelling evidence for the role of the RAS genes in human tumorigenesis, however, came from the observation that identical mutations were found in human tumor specimens. Moreover, specific RAS genes were associated with different tumor types: mutant K- RAS was most often identified in pancreatic, colorectal and lung cancers, H-RAS in bladder

5 carcinomas and N-RAS in lymphoid malignancies and melanomas (Santos et al., 1984, Hirai et al., 1985, Hand et al., 1984, Fujita et al., 1984, Gambke et al., 1984, Bos et al., 1985, Padua et al., 1985). Together, these early studies unveiled the potential significance of mutant RAS in the development of human malignancies.

Since then, oncogenic RAS mutations have been found in 33% of all human tumors and have been shown to play a driving role in tumor initiation, maintenance and malignant conversion (Karnoub et al., 2008). K-RAS is the most frequently mutated RAS gene and is most commonly found in pancreatic ductal adenocarcinoma (PDAC), colorectal tumors, and non-small cell lung carcinomas (NSCLC) (Parwani et al., 2003, Johnson et al., 1993, Rodrigues et al., 1990) (Figure 1.4). Oncogenic K-RAS is required at multiple stages of tumorigenic progression, as the induction or ablation of K-RASD12 in the pancreas of transgenic mouse models results in the initiation or regression of established tumors, respectively (Klimstra et al., 1994, Grippo et al., 2003, Collins et al., 2012). Studies performed in K-RAS-mutant pancreatic and lung cancer cell lines has revealed that K-RAS dependency is intimately linked to epithelial differentiation state, as K-RAS-mutant cells retaining an epitheloid gene signature selectively require K-RAS for cell viability (Singh et al., 2009). These findings suggest that epithelial-to-mesenchymal transition subverts the requirement of K-RAS for its oncogenic potential, possibly due to the acquisition of additional mutations or a shift in oncogenic gene dependencies. Further evidence supporting the role of K-RAS in tumor initiation comes from the identification of K-RAS mutations in pre- neoplastic lesions (Klimstra et al., 1994, Tada et al., 1996). Moreover, siRNA depletion of oncogenic K-RAS reduces the growth and metastases of PDAC xenograft models (Zhu et al., 2006, Fleming et al., 2005). A similar paradigm of early K-RAS oncogene activation and malignant promotion has been observed in colorectal cancer and NSCLC (Vogelstein et al., 1988, van Etten et al., 2002, Westra et al., 1996, Wang et al., 2006). N-RAS mutations, while not as widespread as K-RAS mutations, occur with high frequency in hematologic malignancies and correlate with poor prognosis (Neri et al., 1988, Lubbert et al., 1990) (Figure 1.4). Moreover, N- RAS has been shown to drive the initiation and propagation of tumorigenesis in several malignant myeloid subtypes (Padua et al., 1988, Byrne et al., 1998, Shen et al., 2011, Auewarakul et al., 2006, Emanuel, PD, 2008). By distinction, H-RAS is rarely mutated in human tumors, although cases with alterations in this factor have been found in bladder, kidney, thyroid and breast cancers (Adjei et al., 2001) (Figure 1.4).

Figure 1.4 RAS isoform mutations in human cancer. K-RAS, N-RAS and H-RAS mutations account for 86%, 11% and 3% of total RAS mutations in human cancers, respectively (Downward, J, 2003). K-RAS mutations are most commonly found in PDAC (>95%), colorectal (>50%) and NSCLC (>30%), while N-RAS is frequently mutated in hematologic malignancies, including 20% of acute lymphoblastic leukemias (ALL), 30% of acute myeloid leukemias (AML) and 60% of chronic myelomonocytic leukemias (CMML). In order to be active, both wild-type and mutant forms of RAS must be anchored to the plasma membrane. This is achieved through post-translational modifications in their hypervariable (HV) C-terminii (Figure 1.2). These domains contain a CAAX membrane targeting sequence that is farnesylated by the enzyme farnesyltransferase (FTase) (Schaber et al., 1990). RAS is subsequently transported to the endoplasmic reticulum (ER) where the RAS-converting enzyme 1 (RCE1) recognizes the CAAX motif and cleaves the –AAX sequence, allowing for isoprenylcysteine carboxymethyltransferase 1 (ICMT1) to carboxymethylate the final Cysteine residue (Choy et al., 1999). The K-RAS-4B isoform contains a stretch of lysines that allow its direct interaction with the plasma membrane, while H-, N- and K-RAS-4A must be palmitoylated by palmitoyltransferase (PTase) in order to stabilize this interaction (Hancock et al., 1990). K-RAS-4A and N-RAS may also undergo prenylation by geranylgeranyl transferase (GGTase) in order to secure their association with the plasma membrane (Wright et al., 2006). These numerous posttranslational processes provide the biochemical basis for why the first chemotherapeutic inhibitors of the RAS GTPases, farnestyltransferase inhibitors (FTIs), were ultimately ineffective. Although they efficiently prevented the farnestylation reaction, K-RAS could undergo alternative prenylation reactions that rescued its plasma membrane localization and oncogenic capacity (Whyte et al., 1997, Lobell et al., 2001).

Once at the membrane, wild-type RAS can be activated by a number of upstream regulators, including receptor tyrosine kinases (RTKs), G-protein coupled receptors (GPCRs), integrins and immune receptors. Activation of RAS was first described in response to epidermal growth-factor (EGF) and was found to be a critical mediator of serum-induced mitogenic responses in murine fibroblasts (Kamata et al., 1984, Mulcahy et al., 1985). Later it was found that membrane- associated receptors initiated the GTP-loading of RAS by activating their respective GEFs, including SOS1, SOS2 and RASGRF, or inactivating GAPs, such as p120RASGAP and NF1 GAP, via receptor-associated cytoplasmic proteins like growth factor receptor-bound protein-2 (Grb2) (Molloy et al., 1989, Xu et al., 1990, West et al., 1990, Gale et al., 1993, Li et al., 1993).

Once activated, the RAS GTPases bind a large number of effector molecules and elicit diverse biological signals regulating gene transcription, cell proliferation, differentiation and survival. The most thoroughly studied effectors of RAS include Raf, phosphatidyl inositol-3 kinase (PI3K) and Ral-guanine nucleotide dissociation stimulators (Ral-GDS) (Figure 1.5). The Serine/Threonine (Ser/Thr) kinase Raf was the first RAS effector identified and was established as a critical mediator of RAS-induced mitogenic changes (Moodie et al., 1993, Warne et al., 1993, Zhang et al., 1993, Vojtek et al., 1993). Raf signals through a kinase cascade involving the sequential phosphorylation and activation of mitogen-activated extracellular signal-regulated kinase (MEK1/2 or MAPKK) and extracellular signal-regulated kinase (ERK1/2 or MAPK). Phosphorylation of ERK1/2 can result in the initiation of transcription via its nuclear translocation, or signal transduction via the phosphorylation of its cytoplasmic targets (Leevers and Marshall, 1992). The importance of the Raf/MAPK pathway was first observed when it was found that Raf activation was critical for cellular transformation induced by oncogenic RAS (Khosravi-Far et al., 1995). Overexpression of active Raf was also sufficient to induce transformation in murine fibroblasts (White et al., 1995). Over the years the importance of the MAPK pathway in RAS tumorigenesis has become increasingly clear, as MEK activity is required for tumor growth in RAS-mutated pancreatic, colorectal, lung and breast tumors (Engelman et al., 2008, Hoeflich et al., 2009). The identification of mutually exclusive B-Raf and RAS mutations in human tumors has also emphasized the significance of aberrant Raf/MAPK signaling in human oncogenesis (Rajagopalan et al., 2002).

Figure 1.5 RAS effector pathways. RAS has a number of known effectors, the best studied of which are PI3K, PLC, Raf, TIAM1 and RalGDS. PI3K catalyzes the phosphorylation of phosphatidylinositol (4,5)-bisphosphate (PIP2) to its (3,4,5)-triphosphate form (PIP3), which can activate AKT/PKB to mediate cell survival and protein synthesis through IKK/NF-B and mTOR/S6K pathways, respectively. Phospholipase C epsilon (PLC) cleaves PIP2 to generate diacylglycerol (DAG), which can activate protein kinase C (PKC). TIAM1 and Ral-GDS are Rac and Ral GEFs, respectively, that can activate p21 activated kinases (PAKs) and tank-binding kinase 1 (TBK1). Each of these effectors is required for full transformation downstream of oncogenic RAS. The central arm of the RAS pathway is the mitogenic Raf/MEK/ERK cascade, which is a main determinant of RAS transformation. Shortly after the characterization of the RAS/MAPK cascade, the PI3K and Ral-GDS RAS effectors were identified (Rodriguez-Viciana et al., 1994, Hofer et al., 1994). PI3K activity was shown to be required for RAS transformation in NIH 3T3 cells via its anti-apoptotic effects, mediated by the Ser/Thr kinase AKT/protein kinase B (PKB) and the transcription factor nuclear factor-kappa B (NF-B), both of which are potent regulators of cell survival (Rodriguez-Viciana et al., 1997, Marte et al., 1997, Mayo et al., 1997). AKT can activate NF-B by inactivating its inhibitor IKK through direct and mammalian target of rapamycin (mTOR)-dependent

phosphorylation (Ozes et al., 1999, Dan et al., 2008). However, NF-B can also be activated by Raf and Ral-GDS-induced IKK phosphorylation in response to oncogenic RAS, thereby increasing the complexity of the requirement for NF-B in RAS-mediated cellular transformation (Norris et al., 1999). Like Raf, PI3K is critical for tumorigenesis in several RAS- mutant human tumors, and together they are thought to be the principal contributors to the RAS- transformed phenotype (Campbell et al., 2007, Engelman et al., 2008, Hoeflich et al., 2009). This is supported by recent studies showing synergism in the anti-tumor effects exerted by the combination of MEK and PI3K inhibitors in human PDAC, lung, colorectal and breast cancers (Campbell et al., 2007, Engelman et al., 2008, Yu et al., 2008, Hoeflich et al., 2009 and 2012).

The Ral-GDS proteins, GEFs for RalA and RalB GTPases, contribute to RAS tumorigenesis in human epithelial cell line models and are required for H-RAS-induced skin tumor formation (White et al., 1996, Urano et al., 1996, Gonzalez-Garcia et al., 2005). Many other RAS effectors shown to play functionally important roles downstream of oncogenic RAS include phospholipase C (PLC), T-cell lymphoma invasion and metastasis-1 (TIAM1), Ras interaction/interference protein-1 (RIN1), ALL (acute lymphoblastic leukaemia)-1 fused gene on chromosome 6 (AF-6) and the RAS association domain-containing family (RASSF) proteins (Kelley et al., 2001, Lambert et al., 2002, Kuriyama et al., 1996, Han et al., 1995) (Figure 1.5). The large number of identified targets that contribute to RAS-mediated cellular transformation in vitro and in vivo has led to the paradigm that many downstream effectors of RAS cooperate to induce a fully transformed phenotype.

1.3 The Rho subfamily

The Rho GTPases are critical modulators of cell morphology and motility through their regulation of the actin cytoskeleton. The Rho family is comprised of 22 members in humans and includes Rho (A, B and C isoforms), Rac (1, 2 and 3 isoforms), Cdc42 (G25K and Cdc42Hs isoforms), RhoD, RhoG, TC10, Rnd (1, 3 and 6 isoforms) and TTF (Wennerberg et al., 2005). The best studied members include Rho, Rac, and Cdc42, which were identified by their abilities to generate distinct actin structures in fibroblast cells. Activation of Rho by lysophosphatidic acid (LPA) in serum-starved Swiss 3T3 fibroblasts resulted in the formation of long, parallel

10 bundles of polymerized actin known as stress fibres (Nobes and Hall, 1992). By distinction, platelet-derived growth factor (PDGF) or EGF stimulation of Rac was observed to promote actin-based membrane ruffling and lamellipodia formation at the cell periphery (Nobes and Hall, 1995). Activation of Cdc42 by the GPCR agonist bradykinin, in turn, produced finger-like projections at the front edge of the cell known as filopodia (Kozma et al., 1995). Rho, Rac, and Cdc42 also regulate the formation of focal adhesion complexes, which mediate cell-ECM interactions and function as signaling hubs, connecting extracellular signals to the intracellular space (Nobes and Hall, 1995, Hotchin and Hall, 1995). Together, these effects cooperate to control many actin-based processes including cell motility, cell migration, cytokinesis, phagocytosis, pinocytosis, morphogenesis and axon guidance (Nobes and Hall, 1999, Prokopenko et al., 2000, Cox et al., 1997, Settleman, J, 1999, Luo et al., 1997). Rho GTPase activation has been established as a critical requirement for growth factor-mediated cell migration in fibroblast and epithelial cells (Takahashi et al., 1993 and 1995, Ridley et al., 1995). Moreover, inhibition of Rho GTPases using the C3 toxin from Clostridium botulinum or dominant negative mutants prevents the migration and invasion of tumor cells (Yoshioka et al., 1995, Verschueren et al., 1997, Habets et al., 1994, Keely et al., 1997). Like the RAS GTPases, these early studies were clear indicators of the potential role of Rho GTPases in tumor progression.

The Rho family exhibits important actin-independent functions, including the regulation of gene expression and cell proliferation. Rac1 and Cdc42, and in some cell types RhoA, can activate the c-jun N-terminal kinase (JNK) and p38 MAPKs, which activate ATF-2 and Jun transcription factors (Coso et al., 1995, Minden et al., 1995, Bagrodia et al., 1995a). RhoA, Rac1 and Cdc42 also activate the serum response factor (SRF) and NF-B transcription factors (Hill et al., 1995, Perona et al., 1997). Rho GTPases regulate the transcription of a diverse collection of genes, including those involved in cell cycle progression, apoptosis, cytoskeletal components and the inflammatory response. The role of Rho GTPases in cell-cycle progression was established early on with the elucidation that RhoA, Rac1 and Cdc42 are all required for progression through the G1 phase of the cell-cycle in response to serum stimulation (Yamamoto et al., 1993). C3 treatment of Swiss 3T3 fibroblasts results in the accumulation of cells in G1 and an inhibition of cell growth (Yamamoto et al., 1993). In support of these findings, microinjection of constitutively active mutants of RAS, RhoA, Rac1 and Cdc42 causes G1 progression and

11 stimulation of DNA synthesis in quiescent 3T3 cells, which can be blocked by co-expression of their respective dominant-negative mutants (Olson et al., 1995). The effects of Rho proteins on cell proliferation can largely be attributed to gene expression changes in cell cycle-associated genes including several cyclins (cyclin A, cyclin B1, cyclin D1, cyclin E, cyclin F and cyclin G), cell division cycle (cdc) proteins (cdc20, cdc25c, cdc2a, cdc34 and cdc7), the mitotic spindle regulators Aurora kinase A and B (AurkA and AurkB) and the cell cycle inhibitor p21 (Berenjeno et al., 2007, DeGregori et al., 1995, Westwick et al., 1997, Sahai et al., 2001, Hirai et al., 1997). Transcriptional profiling of cells transformed by oncogenic forms of Rho GTPases (RhoAQ63L, RhoBQ63L and RhoCQ63L) revealed that they induce gene expression changes associated with four main transcription factor networks: c-myc, E2F1, p53 and c-Jun (Berenjeno et al., 2007). Although all of c-myc, E2F1 and c-Jun are all required for RhoQ63L-mediated cellular transformation, c-Jun and E2F1 are uniquely required to regulate the expression of genes associated with cell proliferation, while c-myc regulates the expression of genes involved in RhoQ63L-dependent loss of contact inhibition (Berenjeno et al., 2007).

Mirroring the RAS GTPases, the Rho family is strongly implicated in cellular transformation. Early evidence of this came from the observation that overexpression of constitutively active RhoA, Rac1 and Cdc42 could induce morphological transformation of murine and rodent cells (Qiu et al., 1995a and b, Khosravi-Far et al., 1995, Prendergast et al., 1995, Qiu et al., 1997). In contrast to RAS, however, constitutively activated mutants of Rho have not been identified in human tumors to date (Moscow et al., 1994, Rihet et al., 2001). A more common phenomenon involves the overexpression of Rho family members or aberrant expression or activation of one of their regulators. For example, Rho isoforms RhoA and/or RhoC are overexpressed in malignancies of the breast, ovary, colon, pancreas, lung, liver and brain (Horiuchi et al., 2003, Fritz et al., 1999, Suwa et al., 1998, Fukui et al., 2006, Gou et al., 2011). The overexpression of RhoC correlates with advanced tumor grade and decreased survival in serous ovarian carcinoma (SOC), colorectal cancer, pancreatic cancer and, most prominently, breast cancer (Horiuchi et al., 2003, Fritz et al., 1999, Suwa et al., 1998, Yuan et al., 2007). Importantly, the intratumoral injection of RhoA or RhoC siRNAs was shown to inhibit the growth and metastases of xenografted breast cancer cells, demonstrating the functional significance of their upregulation (Pille et al., 2005).

RhoA is associated with tumor progression in ovarian cancer, breast cancer, SCLC, HCC and astrocytomas by mediating their increased migratory, invasive and/or metastastic phenotypes (Horiuchi et al., 2003, Fritz et al., 1999, Varker et al., 2003, Fukui et al., 2006, Li et al., 2006). The Rac GTPases are also dysregulated in human malignancies, however, their increased activity is more commonly associated with the miss-expression of their respective GEFs, GAPs or GDIs. For example, expression of the Rac GEF DOCK180 correlates with advanced tumor grade in ovarian carcinoma (OC) and its silencing prevents the proliferation, motility and invasion of SK- OV-3 OC cells (Zhao et al., 2011, Wang et al., 2010). The Rac GDI RhoGDI12 was found to antagonize the growth, invasion and in vivo lung metastases of OC cells via the inactivation of Rac1, providing further evidence of the importance of Rac activation for ovarian tumor progression (Stevens et al., 2011). The Rac GEF TIAM1, originally found to mediate RAS- induced skin carcinogenesis in mice, has also been shown to promote the migratory and invasive properties of prostate, HCC, PDAC, colon and breast tumor cells (Malliri et al., 2002, Engers et al., 2006, Chen et al., 2012, Cruz-Monserrate et al., 2008, Minard et al., 2006, Strumane et al., 2009). To add to this expanding list are aberrations in expression of the Rac GEFs Vav1-3 and P- Rex1 found in breast, lung and melanoma carcinomas, respectively, which exert their transforming properties via the activation of Rac1 (Citterio et al., 2012, Lazer et al., 2009, Lindsay et al., 2011).

Interestingly, there is a strong connection between RAS and Rho GTPases in cellular transformation. RAS and Rho cross-talk was initially predicted based on the observation that oncogenic RAS induces changes in stress fibres, lamellipodia and filopodia in fibroblast cells (Bar Sagi and Feramisco, 1986). Subsequently, an inhibitory interaction between the RAS GAP p120RASGAP and the Rho GAP p190RhoGAP was identified that resulted in increased RhoA activity in RAS-transformed cells (Settleman et al., 1992). It wasn’t until microinjection studies performed by Hall and colleagues using dominant-negative Rho GTPase mutants, however, that it was established that Rho GTPases play a functional role downstream of oncogenic RAS (Ridley and Hall, 1992). They showed that inhibition of RhoA or Rac1 blocked RASV12-induced stress fibre and membrane ruffle formation, respectively. Importantly, inhibition of RhoA, RhoB, Rac1, Cdc42 or RhoG resulted in a significant reduction in the focus-forming ability of oncogenic RAS (Ridley and Hall, 1992, Prendergast et al., 1995, Khosravi-Far et al., 1995, Lebowitz et al., 1997). Conversely, overexpression of active mutants of RhoA, Rac1 and Cdc42

13 showed cooperative and synergistic focus-forming activity with active Raf1, suggesting that Rho and RAS GTPases work together to promote the transformed phenotype (Khosravi-Far et al., 1995, Whitehead et al., 1998).

The connection between RAS and Rho GTPases is conserved in human tumors expressing endogenous mutations in RAS. In MCF-7 breast and HT-1080 fibrosarcoma cells, migration induced by EGF or constitutively active MEK1 is prevented by dominant-negative RhoA or chemical inhibition of its downstream effector ROCK (Jo et al., 2002). Here, inhibition of RhoA failed to prevent the migratory phenotype induced by ERK-independent factors, suggesting that RhoA is specifically activated by the RAS/MAPK pathway to promote tumor progression. Moreover, upregulation of the RAS target TrkBT1, which sequesters a RhoGDI, promoted the metastasis of PDAC cells by increasing RhoA activation (Li et al., 2009). Both B-Raf and RAS oncogenes have been shown to regulate Rho GTPase pathways to induce migration and invasion of human colon cancer cells (Makrodouli et al., 2011). The Rac1 GTPases have also been implicated downstream of oncogenic RAS in human tumors. A classic example of this is in the case of TIAM1, which is required for H-RAS-induced skin tumor growth but promotes the metastatic conversion of established tumors (Malliri et al., 2002). Further studies showed that TIAM1 is a direct effector of RAS via a bona fide RAS binding domain (RBD) in its N-terminus (Lambert et al., 2002) (Figure 1.6). Mouse models of adenomatous polyposis coli (APC)-induced colon cancer and ERBB2-induced mammary cancer parallel the dichotomy of TIAM1 function in H-RAS-induced skin tumors, with reduced primary tumor formation but increased metastases in the absence of TIAM1 (Malliri et al., 2006, Strumane et al., 2009). Its metastasis-suppressing function is further supported by the inverse correlation between malignant progression and TIAM1 protein expression in breast cancer (Stebel et al., 2009). These studies highlight the complexity and evolution of the signaling interplay between RAS and Rho GTPases in different contexts and suggest that the requirement of Rho GTPases in RAS-mediated cellular transformation may depend on different stages of tumorigenic progression.

In contrast to the Rac family, the mechanisms connecting RAS to Rho remain largely unclear. As the Rho GEFs are known to contribute to cancer progression, as I will describe in the next section, it has long been thought that RAS may activate Rho GTPases via the regulation of Rho

GEFs. However, as of yet there is a lack of evidence to support this hypothesis; thus, the search for mediators of RAS and Rho cross-talk continues to be an area of intense study.

1.4 Guanine nucleotide exchange factors

Guanine nucleotide exchange factors (GEFs) are critical regulators of GTPase activation in mammalian cells. A striking feature of GEFs is that they outnumber their GTPase substrates by a factor of 3 (Venter et al., 2001). This is exemplified by the mammalian Rho family, which consists of 83 GEFs and only 22 GTPases. Although the reasons for this redundancy have not been fully determined, it is thought that distinct GEFs dictate the precise cellular function of the otherwise pleiotropic GTPases by connecting them with different upstream receptors and/or signaling molecules in the cell.

The first identified GEF, the Rho GEF Dbl, was isolated as a transforming protein in NIH 3T3 fibroblasts derived from DNA from a diffuse B-cell lymphoma (Eva and Aaronson, 1985). It was found to display significant homology to an activator of the GTPase Cdc42, Cdc24, and was subsequently also shown to catalyze association with GTP (Ron et al., 1991, Hart et al., 1991). The conserved domain shared by Cdc24 and Dbl was termed the Dbl homology (DH) domain and is the site of catalytic exchange (Hart et al., 1994). Although DH domains are a structurally conserved component of all GEFs, they display surprisingly low sequence homology (20%) (Hart et al., 1994). However, their three dimensional structures are highly similar and consist of 11 alpha helices, two of which are exposed on the surface of the protein and participate in the formation of the GTPase-interacting pocket (Cherfils and Chardin, 1999). GEFs activate their substrates by binding to the GDP-bound form of GTPases and destabilizing the GDP-GTPase interaction, favouring the formation of a nucleotide-free intermediate. Since the approximate ratio of GTP to GDP at physiological conditions is 10:1, with GTP concentrations ranging from 200-500M, the binding of GTP and activation of the GTPase is favored (Traut, TW, 1994, Cherfils and Chardin, 1999). A pleckstrin homology (PH) domain is typically located adjacent and C-terminal to the DH domain and together they form the core catalytic module and minimal structural unit required to promote in vivo nucleotide exchange. PH domains in other signaling molecules are capable of binding phosphatidylinositol phospholipids (PIPs) and are thus

15 classically thought to function by mediating plasma membrane interactions (Rebecchi et al., 1998, Lemmon et al., 2000). However, in the case of many GEFs the contribution of the PH domain is multifaceted. Only in some GEFs can the PH domain be functionally replaced by a membrane-targeting phospholipid; moreover, other studies have shown that the PH domain can directly affect the catalytic activity of the DH domain by mediating inhibitory protein interactions or through intramolecular binding (Figure 1.6).

Figure 1.6 Crystal structure of the DH-PH domain of Dbl’s big sister (Dbs) in complex with RhoA. The DH (depicted in blue) and PH (depicted in yellow) domains of the Rho GEF Dbs contribute to the binding of the RhoA GTPase (depicted in green). The 3 and 4 loops of the DH domain and the 6 helix of the PH domain form significant interactions with RhoA and stabilize the conformation of the catalytic core. N designates N-terminal; C designates C-terminal;  designates alpha;  designates beta (Worthylake et al., 2004). In contrast to their shared catalytic domains, there is considerable structural diversity among GEFs for a specific GTPase (Figure 1.7). This is especially striking for the Rho GEFs, which contain a plethora of additional functional regions such as Src homology 2 and 3 (SH2 and SH3) domains, phospholipid binding motifs, Ras-GEF domains, coiled-coil regions, cysteine-rich domains, zinc finger binding motifs, Rho-GAP domains and PDZ and RGS domains, among others. These flanking regions confer specificity in protein-protein or protein-lipid interactions, second messenger binding and protein kinase phosphorylation in response to upstream stimuli, thus mediating the selectivity of the GTPase response. These regions can also promote proper localization or have autoregulatory functions, thereby regulating the spatial or temporal

16 activation of Rho GTPases, respectively. Moreover, the multitude of additional Rho GEF domains suggests that GEFs may have functions independent of Rho GTPase activation.

Figure 1.7 The diversity in Rho GEF domain organization. In addition to the core DH and PH catalytic module, Rho GEFs contain many other domains that are subject to diverse regulatory interactions. Vav is activated by src via a consensus phospho-Tyrosine motif and contains additional src homology 2 and 3 (SH2 and SH3), calponin- homology (CH) and Cysteine-rich (CR) domains. Arhgef12 (also known as Leukemia-Associated Rho GEF (LARG) and KIAA0383) contains a regulator of G protein signaling (RGS) domain that is crucial for its activation downstream of G protein coupled receptors (GPCRs) via the G family of heterotrimeric GTPases. APC influences Arhgef4 (also known as APC-stimulated GEF (ASEF), STM6 and KIAA1112) migration and adhesion by binding to a conserved APC binding region (ABR) in its N-terminus. TIAM1 is directly activated by RAS via a bona fide RAS binding domain (RBD) and contains an additional N-terminal PH domain. TIAM1 and Arhgef12 also contain PDZ domains, which mediate a wide spectrum of protein-protein interactions. Evidence for the autoregulatory role of the sequences flanking the core DH-PH domains of Rho GEFs was apparent in early studies showing that their truncated mutants exhibited increased transforming capacity in focus-forming assays (Whitehead et al., 1997). Removal of the N- terminal sequences of Dbl, Vav, Asef, TIAM1, Ect2 and Net1 or the C-terminal sequences of p115RhoGEF and AKAPLbc resulted in their constitutive activation (Eva et al., 1985, Toksoz and Williams, 1994, Chan et al., 1996, Whitehead et al., 1995b, Miki et al., 1993, Chan et al., 1994). In most of these cases, genomic deletion of coding sequences was initiated by the transfection procedure as opposed to genetic events in the cancer cells themselves; however, it revealed their potential as oncogenes. The mechanism underlying the auto-inhibition exhibited by some Rho GEFs is best characterized in the case of Vav, in which an N-terminal Tyrosine (Tyr174) interacts directly with the DH domain and blocks its interaction with GTPases. Upon

17 receptor stimulation, Tyr174 is phosphorylated by src, inducing a conformational change in the N-terminus and relieving steric hindrance of the DH domain (Salojin et al., 1999, Lopez-Lago et al., 2000). In contrast, the N-terminus of Dbl interacts with its PH domain via heat shock cognate protein 70 (Hsc70), thereby physically preventing GTPase substrate binding (Ron et al., 1989, Bi et al., 2001, Kauppinen et al., 2005). Moreover, Hsp90 interacts with Hsc70 and the N-terminus of proto-Dbl and induces its ubiquitination, such that deletion of this region results in both its accumulation to high levels in the cell and its constitutive GEF activity. These mechanistic studies elegantly account for the potent transforming activity of oncogenic Dbl and provide insight into the diverse modes of Rho GEF regulation (Kamynina et al., 2007).

The importance of Rho GTPase-regulated pathways in cancer is highlighted by the identification of genetic alterations in many Rho GEFs in human malignancies and complements the observation that mutations in the Rho GTPases themselves are rare events. Indeed, growing evidence supports a critical role for Rho GEFs in the dysregulation of GTPase signaling in human cancers (described below). Aberrations in Rho GEFs found in human cancers include rearrangements, deletions, overexpressions and mutations that confer increased catalytic activity to the proteins. However, in some instances the contribution of Rho GEFs to tumorigenesis is independent of its enzymatic activity, adding another layer of complexity to Rho GEF function in human tumorigenesis.

Several Rac GEFs have been implicated in tumorigenesis, including TIAM-1, Vav1-3 and P- Rex-1 and 2a. TIAM-1 is overexpressed in a number of human malignancies and is a direct target of H-RAS (Chen et al., 2012, Lambert et al., 2002). TIAM-1 is required for the initiation of tumorigenesis but contributes to the metastatic conversion of established tumors (Malliri et al., 2002). The Vav1 Rac GEF is overexpressed in PDAC cells as a result of promoter demethylation and is required to support anchorage-independent growth and xenograft growth in vivo; similar observations have also been made in lung cancer cells (Fernandez-Zapico et al., 2005, Lazer et al., 2009). Vav2 is hyperactivated in response to EGFR signaling and mediates invasion of tumor cells in head-and-neck squamous cell carcinoma (HNSCC) (Patel et al., 2007). Vav3 is overexpressed in glioblastoma, androgen-independent prostate cancer cell lines, and breast tumors and contributes to the invasion of glioblastoma cells (Salhia et al., 2008, Lyons et al., 2006, Lee et al., 2008). The phosphatidylinositol-3,4,5-triphosphate-dependent Rac exchange

18 factor 1 (P-Rex1) is overexpressed in metastatic prostate cancer patient samples and cell lines and mediates the migration and invasiveness of these cells via the activation of Rac1 (Qin et al., 2009). Interestingly, P-Rex2a was found to contribute to breast cancer progression by directly interacting with and inhibiting the PTEN tumor suppressor independently of its GEF activity (Fine et al., 2009). These studies provide evidence that Rho GEFs can modulate oncogenic pathways in both GEF-dependent and independent manners.

In addition to being overexpressed or aberrantly activated by upstream regulators, Rho GEFs are structurally altered in many human cancers. The BCR-ABL translocation is famous for its well- described rearrangement in Philadelphia chromosome-positive leukemias, which comprise 90% of chronic myelogenous leukemias (CMLs) (Heisterkamp et al., 1985, Laurent et al., 2001). The 9:22 chromosomal translocation results in the fusion of the N-terminal regulatory sequences of the Rho GEF breakpoint cluster region (BCR) protein with the kinase domain of the non-receptor tyrosine kinase ABL. Although BCR contains both a Rho GEF and Rho GAP domain, only the Rho GEF domain is present in the chimera. The oncogenic capacity of the fusion protein is predominantly mediated by the constitutive activation of ABL, however, it displays a partial dependence on RhoA nucleotide exchange for anchorage-independent growth (Laurent et al., 2001). The Rho GEF LARG was also identified as a rearranged gene with the mixed-lineage leukemia (MLL) gene in a patient with acute myeloid leukemia (AML) (Kourlas et al., 2000). The fusion protein retains the DH-PH catalytic core and loses the N-terminal auto-inhibitory region of LARG, but whether it exhibits constitutive RhoA exchange in AML remains unclear. Interestingly, while MLL-associated LARG acts as an oncogene, wild-type LARG has been implicated as a tumor suppressor in human breast and colorectal cancer (Ong et al., 2009). Thus, the presence of the PDZ domain in the N-terminus of LARG may mediate GEF-independent, tumor-suppressive interactions. Another study, however, found that LARG induced foci formation in fibroblasts but decreased invasion in lymphoma cells, both in GEF-dependent manners, suggesting that wild-type LARG mirrors TIAM1 in its opposing roles in the initiation and progression of cellular transformation (Jiang et al., 2010). These studies reinforce the need to study the role of Rho GEFs at multiple stages of tumorigenic progression and in response to diverse stimuli in order to gain a full understanding of their contribution to tumorigenesis.

Together, the mechanistically varied regulation of Rho GEFs and the evidence being compiled concerning their functional roles across all tumor types mounts a strong argument for them as a class of potent human oncogenes. Moreover, the reasons underlying the imbalance between the number Rho GEFs and their catalytic substrates is becoming more obvious with the elucidation of novel GEF-independent functions and as we learn more about the diversity in their upstream regulators. With only a fraction of Rho GEFs studied in the context of cancer, however, it remains to be established whether the remaining dysregulated GEFs in human tumors are passengers or drivers of oncogenesis.

1.5 Arhgef2

The Rho GEF Arhgef2, also known as murine Lfc or human GEF-H1, is a guanine nucleotide exchange factor for RhoA (Krendel et al., 2002). It was originally identified in a C-terminally truncated form whose overexpression induced morphological transformation of NIH 3T3 cells (Whitehead et al., 1995). An N-terminal mutant was also found to initiate tumor formation in mouse xenograft models, supporting the paradigm that sequences surrounding the core DH-PH module can exert a GEF-inhibiting effect (Brecht et al., 2004). Arhgef2 contains an N-terminal C1, or zinc-finger, domain and two C-terminal coiled-coil motifs as well as several Ser/Thr phosphorylation sites, which contribute to its regulation and function through a multitude of established binding partners (Figure 1.8).

Figure 1.8 The domain organization of Arhgef2. Arhgef2 contains an N-terminal cystein rich (C1) domain and two C-terminal coiled-coil (CC) domains that flank the DH-PH catalytic unit. Glutamate (E) and Lysine (K) amino acids at residues 243 and 394, respectively, are critical for catalytic exchange. Serine 885 (S885) is a major negative

regulatory site that can be phosphorylated by AurA and B, PAK1 and 4, PKA and Cdk1, resulting in 14-3-3 binding and Arhgef2 inhibition. A number of additional binding partners of Arhgef2 have been identified that functionally interact with the CC, PH and N-terminus of Arhgef2, as indicated above.

Arhgef2 is unique among GEFs in that it associates with and regulates the microtubule array as well as the actin cytoskeleton (Krendel et al., 2002). p190RhoGEF is currently the only other of the 88 Rho GEFs that has been shown to bind microtubules (van Horck et al., 2001). Microtubule dynamics are tightly coupled to the actin cytoskeleton during cell migration, where microtubule depolymerisation and subsequent formation of actin stress fibers and actomyosin- dependent cell contractility allows cells to dynamically propel themselves forward (Nobes and Hall, 1992). Although the Rac and Rho GTPases have long been known to mediate the succession in morphological changes linking these two cytoskeletal networks, neither GTPase has shown localization to the microtubule array. Thus, the identification of Arhgef2’s association with microtubules suggested a novel mechanism by which microtubule depolymerisation may be linked to Rho GTPase-dependent actin stress fiber formation. A critical study by Krendel et al. revealed that Arhgef2 interacts with microtubules via its N- and C-terminal ends, resulting in the inhibition of its GEF activity toward RhoA (Krendel et al., 2002). Overexpression of full-length Arhgef2 results in microtubule bundling and increased resistance against the microtubule depolymerising agent nocodazole, demonstrating that Arhgef2 stabilizes the microtubule array. Furthermore, deletion of N- and C-terminal portions of Arhgef2 results in its translocation from microtubules to the actin cytoskeleton, where it initiates the formation of stress fibers in a RhoA- dependent manner. Interestingly, although microtubule-associated Arhgef2 displays weak exchange activity compared to the cytoplasmically-localized deletion mutants in cells, their in vitro exchange activities are similar, showing that their inhibitory properties are conferred by in vivo protein-protein or microtubule-dependent interactions rather than auto-inhibitory functions. Indeed, recent work in our laboratory has shown that the inhibition of Arhgef2 is mediated through T-complex testis-specific protein-1 (Tctex-1), a light chain of the dynein motor complex that functions in microtubule transport (Meiri et al., 2012). Tctex-1 binds to amino acids 87-151 of Arhgef2, thereby linking Arhgef2 to the microtubule array and enabling its inhibition (Figure 1.8) (Meiri et al., 2012). Importantly, deletion of the Tctex-1 binding domain (Arhgef287-151) results in the release of Arhgef2 from microtubules, elevated GEF activity in the cytoplasm and increased stress fiber formation. Together, these studies consolidate Arhgef2 as the first known mediator connecting microtubule dynamics to actin polymerization.

Given its role as a critical regulator of microtubules and the actin cytoskeleton, it comes as no surprise that Arhgef2 has been implicated in biological processes involving the maintenance of cell structure, polarity, motility, migration and cell cycle progression. The first study looking at Arhgef2 function in epithelial cells found that Arhgef2 is a tight junction-associated protein (Benais-Pont et al., 2003). Tight junctions are one of four main junction types that link epithelial cells together in order to form a compact epithelium, required to line and protect the organs of the body. Tight junctions are the most apical of the intercellular junctions and regulate selective paracellular diffusion and restrict the intermixing of apical and basolateral membrane components, thereby maintaining cell polarity (Cereijido et al., 2000). Benais-Pont et al. found that Arhgef2 is associated with tight junctions in interphase cells and promotes RhoA-dependent increases in the paracellular permeability of small molecular weight proteins (Benais-Pont et al., 2003). Later studies showed that Arhgef2 is recruited to tight junctions by the adaptor cingulin, which mediates its inhibition in confluent cells (Aijaz et al., 2005). Moreover, cingulin depletion results in the release of Arhgef2 from tight junctions, RhoA activation, and increased cell proliferation (Aijaz et al., 2005). Later, members of the same laboratory found that Arhgef2 could also localize to apical junctions via a similar junctional adaptor protein, paracingulin, which also resulted in a decrease of its GEF activity at confluency (Guillemot et al., 2008). In calcium-depleted colonic epithelial cells, Arhgef2 activation induces the disassembly of the epithelial barrier by disrupting apical junction complexes through the formation of contractile actomyosin structures in RhoA-ROCK-dependent manner (Samarin et al., 2007). Since calcium levels regulate microtubule dynamics, it was hypothesized that destabilization of the microtubule array and subsequent release of Arhgef2 into the cytoplasm was the triggering event for RhoA- mediated apical junctional dissolution (Samarin et al., 2007).

A role for Arhgef2 has also been established in endothelial cells, which form a semiselective permeable barrier between the blood and interstitial space and regulate macromolecule and leukocyte transport through the vessel wall. Birukova and colleagues found that depletion of Arhgef2 or expression of dominant-negative mutants of Arhgef2 significantly attenuated thrombin and nocodazole-induced vascular permeability increases and RhoA-mediated cellular events (Birukova et al., 2005). Moreover, they later showed that Arhgef2 mediates increases in vascular endothelial permeability associated with acute lung injury, thereby extending its importance in endothelial barrier function to a disease context (Birukova et al., 2010).

Arhgef2 function is exploited by the bacteria Enteropathogenic Escherichia coli (EPEC), which exerts its pathogenicity by inducing microtubule disruption and Arhgef2 activation, resulting in increased paracellular permeability and degeneration of the colonic epithelium (Caron et al., 2006, Matsuzawa et al., 2004). Arhgef2 is also activated in response to Shigella bacterial invasion in the intestinal epithelium, where it enables cell entry and the activation of the innate immune response via a RhoA-NFB pathway (Fukazawa et al., 2008). Arhgef2 mediates increased kidney tubular epithelial cell permeability in response to the pro-inflammatory cytokine TNF, implicating Arhgef2 in the disruption of tubular cell integrity associated with kidney injury (Kakiashvili et al., 2009, Kakiashvili et al., 2011). Together, these studies demonstrate an important role for Arhgef2 in the regulation of junctional integrity in multiple epithelial cell types and disease contexts. Moreover, they suggest that Arhgef2 may play a role in other diseases involving the abrogation of proper epithelial cell structure, such as cancer.

Arhgef2 also regulates cell cycle progression, as was initially demonstrated by Westwick et al. who showed that C-terminally truncated Arhgef2 can induce cyclin D1 expression in NIH 3T3 cells (Westwick et al., 1998). In MCDK cells, depletion of the Arhgef2 inhibitor cingulin results in the release of Arhgef2 from tight junctions and progression through G1 of the cell cycle (Aijaz et al., 2005). The requirement of Arhgef2 for progression at multiple stages of mitosis has since been shown in fibroblasts and epithelial cells (Bakal et al., 2005, Birkenfeld et al., 2007). Arhgef2 is required for pro-metaphase/metaphase transition in Rat-2 cells (Bakal et al., 2005) and for the localized activation of RhoA at the cleavage furrow during cytokinesis (Birkenfeld et al., 2007). Moreover, Arhgef2 is a phosphorylation target of ERK1/2 and mediates cell proliferation in response to phorbol 12-myristate 13-acetate (PMA) stimulation (Fujishiro et al., 2008).

Early studies showing that Arhgef2 mutants deficient in microtubule binding induced stress fibers and cell contractility were indicators of its potential role in the coordination of cell migration (Krendel et al., 2002). Indeed, endogenous Arhgef2 was later shown to be required for nocodazole-induced increases in actomyosin contractility in HeLa cells (Chang et al., 2008). Elegant studies by Nalbant et al. using fluorescence resonance energy transfer (FRET) biosensors to determine the spatial distribution of RhoA activity in migrating cells showed that depletion of Arhgef2 suppresses RhoA activation at the leading edge, resulting in their decreased

23 migratory capacity (Nalbant et al., 2009). By contrast, studies by Heasman et al. in T cells during transendothelial migration showed that Arhgef2 regulates the actomyosin-based contraction of the uropod in the rear of the cell but has no effect on RhoA activation at the leading edge (Heasman et al., 2010). These reports suggest that Arhgef2 activates distinct cellular pools of RhoA in migrating cells and can regulate both their forward protrusion or tail retraction, depending on cell type and/or context. Arhgef2 also affects focal adhesion turnover in migrating cells and contributes to increased cell rigidity in response to integrin-mediated focal adhesion kinase (FAK)/RAS/ERK activation (Nalbant et al., 2009, Guiluy et al., 2011).

The role of Arhgef2 in cell migration and attachment suggest that Arhgef2 may play a role in the migratory and invasive properties of cancer cells. Several recent reports have demonstrated that Arhgef2 contributes to the invasion and in vivo metastases of breast cancer cells and the migration of HCC cells (Liao et al., 2012, Cheng et al., 2012). Arhgef2 is transcriptionally upregulated in metastatic breast cancer cells by the oncogenic transcription factor hPTTG1 and is activated by Heparanase in brain metastatic breast cancer (BMBC) cells (Liao et al., 2012, Ridgway et al., 2012). There, the upregulation and activation of Arhgef2 resulted in increased breast cancer cell metastases and transmigration through the blood-brain barrier, respectively, demonstrating the functional significance of Arhgef2 dysregulation. Arhgef2 was also identified as an irradiation-responsive gene in breast cancer cells harboring BRCA1/2 mutations, suggesting that Arhgef2 expression may be a potential therapeutic marker in breast cancer (Walker et al., 2008). In HCC, Arhgef2 undergoes genomic amplification, leading to increased expression and increased cell migration (Cheng et al., 2012). Moreover, Arhgef2 was shown to be transcriptionally upregulated by gain-of-function (GOF) mutants of p53 in NSCLC, thereby contributing to their increased proliferative capacity (Mizuarai et al., 2006). Together, these studies suggest that transformed cells can select for increased Arhgef2 expression or activity by distinct mechanisms to promote tumor progression.

The aforementioned studies reveal critical functions for Arhgef2 as a regulator of cell morphology, epithelial cell integrity, cell-cycle progression, migration and adhesion. These functions are mediated via the regulated activation of RhoA and are intimately related to its association to and release from the microtubule array. The important physiological activities of Arhgef2 suggest that its dysregulation may contribute to tumorigenesis. Current research is

24 directed at addressing this question, as evidenced by recently published studies implicating Arhgef2 in the progression and poor prognosis of breast and hepatocellular carcinoma (Liao et al., 2012, Cheng et al., 2012).

In 2000, a report in Nature Genetics looked at the genome-wide transcriptional changes induced by oncogenic H-RAS in rat fibroblast cells (Zuber et al., 2000). Arhgef2 was among several hundred genes that were significantly upregulated in H-RASV12-transformed cells compared to their wild-type counterparts. I read this paper when I joined the lab in 2006 and immediately sought to question the potential role of Arhgef2 downstream of oncogenic RAS. I hypothesized that Arhgef2 may be the Rho GEF linking increased RAS activity to elevated levels of active RhoA in tumor cells. Moreover, I predicted that Arhgef2 may contribute to the malignant conversion of RAS-mutated cells via its dual role in epithelial cell junction formation and cell migration. In the pages that follow, I will lead you through my six year quest to determine the role of the Rho GEF Arhgef2 in RAS-induced tumorigenesis. I hope that you find my discoveries as intriguing and exciting as I have experienced them to be at each stage of my scientific journey.

Chapter 2 Arhgef2 Provides a Positive Feedback Loop Required for Signaling Through the Oncogenic RAS Pathway

2.1 Abstract

Activating mutations in RAS are one of the most common oncogenic events in human cancers, however, as of yet they have proven to be pharmacologically intractable targets. Thus, the identification of RAS effectors essential for tumor cell survival is critical to improve treatment strategies in RAS-mutated malignancies. In this chapter, we find that ARHGEF2 is a transcriptional target of the RAS/MAPK pathway. Increased protein expression of Arhgef2 in RASV12-transformed fibroblast cells contributes to cell proliferation, survival, and transformation in vitro and in vivo xenograft models. Moreover, we find that Arhgef2 is required for the activation of the MAPK pathway in response to oncogenic RAS. Importantly, this effect is independent of its Rho GEF activity and instead relies on its novel function as an adaptor protein for a molecular scaffold of the MAPK pathway, Kinase suppressor of RAS-1 (KSR-1). Arhgef2 facilitates the dephosphorylation of KSR-1 on a critical negative regulatory site, Ser392, by recruiting the B’ subunit of protein phosphatase 2A (PP2A) to the KSR-1/MAPK complex. Depletion of Arhgef2 prevents RASV12-mediated MEK1/2 and ERK1/2 activation in a manner that depends on KSR-1 dephosphorylation on Ser392. Together, these data place Arhgef2 in a positive feedback loop where MAPK-dependent increases in Arhgef2 expression potentiate MAPK signaling in RAS-transformed cells. These findings provide insight into mechanisms underlying oncogenic RAS-mediated cell proliferation and survival and highlight the potential of Arhgef2 as a therapeutic target in RAS-mutated cancers.

2.2 Introduction

Due to the high frequency of RAS mutations in human cancer, the signaling pathways regulated by the RAS oncogenes (H-, K- and N-RAS) have been the subject of intense research (Thomas et al., 2007). The RAS GTPases regulate diverse biological processes, including transcription, translation, cell-cycle progression, apoptosis and cell survival (Macara et al., 1996). The specificity of RAS signaling is determined by its interaction with over a dozen downstream effector molecules, the most well-studied being Raf, PI3K and Ral-GDS (Vojtek et al., 1993, Kodaki et al., 1994, Kikuchi et al., 1994). The Raf Ser/Thr kinases (A-Raf, B-Raf and c-Raf) were the first RAS effectors discovered and are major mediators of cell proliferation and survival via the activation of the MAPK cascade, involving the sequential phosphorylations of MEK1/2 and ERK1/2 (Moodie et al., 1993, Warne et al., 1993, Zhang et al., 1993, Vojtek et al., 1993, Galmiche et al., 2010).

The Kinase Suppressor of RAS (KSR-1) was identified in genetic screens in Drosophila and C. elegans designed to isolate mutations in genes that modified the signaling efficiency of oncogenic RAS (Kornfeld et al., 1995, Therrien et al., 1995, Sundaram et al., 1995). Subsequent studies showed that KSR-1 acts as a molecular scaffold to facilitate signal transmission through the Raf/MAPK cascade (Therrien et al., 1996, Michaud et al., 1997, Cacace et al., 1999, Morrison, 2001). KSR-1 is constitutively associated with MEK1/2 and interacts with ERK1/2 and Raf in response to RAS activation (Therrien et al., 1996, Michaud et al., 1997, Cacace et al., 1999). KSR-1 was shown to be required for RASV12-mediated ERK1/2 activation and cellular transformation in mammalian cells in vitro and in vivo (Kortum et al., 2004, Joneson et al., 1998, Razidlo et al., 2004, Nguyen et al., 2002, Lozano et al., 2003, Xing et al., 2003). Consistent with its role as a scaffolding protein, KSR-1 function must be tightly regulated in order to ensure optimal MAPK signaling downstream of RAS activation. In quiescent cells, KSR-1 is phosphorylated on S297 and S392 and held inhibited in the cytosol by 14-3-3 proteins (Ory et al., 2003). Upon RAS activation, KSR-1 is dephosphorylated at S392, the major 14-3-3 binding site, and translocates to the plasma membrane where it can interact with Raf and ERK1/2 to facilitate signal transduction (Ory et al., 2003).

Work by Ory et al. identified the protein phosphatase 2A (PP2A) as the critical phosphatase required for dephosphorylation of KSR-1 on S392 in response to activated RAS (Ory et al., 2003). The requirement of PP2A for KSR-1 function was supported by early genetic studies in Drosophila and C. elegans showing that mutations in PP2A phenocopied a loss of KSR-1 function in a RAS-mutated background (Wassarman et al., 1996, Sieburth et al., 1999). PP2A is a heterotrimeric S/T protein phosphatase composed of a catalytic (C), structural (A) and regulatory (B) subunit. The catalytic and structural subunits constitutively interact to form a core complex to which one of many B subunits can bind (Janssens et al., 2001). Four families of B subunits exist in mammals (B, B’, B’’ and B’’’) that determine the localization and substrate specificity of the holoenzyme (Janssens et al., 2001). While the A and C subunits constitutively associate with KSR-1, the B subunit is induced only upon RAS activation (Ory et al., 2003). However, the mechanism underlying the recruitment of the B subunit to the KSR-1/PP2A A + C holoenzyme complex has not been elucidated.

Arhgef2 has been implicated in tumorigenesis since its discovery, when it was isolated as a transforming protein in NIH 3T3 cells when overexpressed (Whitehead et al., 1995). An N- terminal truncation mutant of Arhgef2 was also shown to induce tumor formation in nude mice (Brecht et al., 2004). Arhgef2 is transcriptionally upregulated downstream of multiple oncogenes, including gain-of-function mutants of p53, the metastasis-associated gene hPTTG1, TGF, oncogenic RAS, and was recently identified as an amplified gene in hepatocellular carcinoma (Mizuarai et al., 2006, Liao et al., 2012, Tsapara et al., 2010, Zuber et al., 2000, Cheng et al., 2012). Arhgef2 was shown to mediate mutant p53-induced cell proliferation and hPTTG1 and TGF-induced cell migration and motility via activation of its downstream effector RhoA (Mizuarai et al., 2006, Cheng et al., 2012, Tsapara et al., 2010). The upregulation of Arhgef2 downstream of oncogenic RAS, however, has not been validated nor has its functional role downstream of RAS been investigated. Thus, we hypothesized that Arhgef2 may link increased RAS signaling to RhoA activation, thereby potentiating the oncogenic potential of RAS-mutated cells.

In this chapter, we confirm that ARHGEF2 is a transcriptional target of the RAS/MAPK pathway and contributes to cell survival and transformation in RAS-transformed cells both in vitro and in vivo xenograft models of RAS tumorigenesis. Importantly, we find that Arhgef2

28 contributes to RASV12-mediated survival and proliferation in a GEF-independent manner. We also uncover a novel role for Arhgef2 as an adaptor protein, linking the B’ subunit of PP2A to KSR-1, thereby promoting the activating dephosphorylation of KSR-1 on S392 and potentiating MAPK signaling in RAS-transformed cells.

2.3 Experimental Procedures

Derivation of ARHGEF2 knockout mice: A targeting construct was designed to insert a loxP site upstream of exon 2, and a loxP-flanked neomycin resistance cassette (in reverse orientation) downstream of exon 2 of the ARHGEF2 gene. The construct was electroporated into the E14K embryonic stem cell (ES) cell line. Correctly targeted ES cells were injected into recipient blastocysts and chimeric mice were bred to C57BL/6 females to establish the colony. The ARHGEF2 floxed mice were then bred with CMV-Cre mice. The resulting mice lacking both exon 2 and the floxed neomycin cassette were selectively bred to remove the CMV-Cre transgene. Heterozygous mice were backcrossed for at least 4 generations and then bred together to generate homozygous mice.

Cell lines and cell culture: MEFs derived from ARHGEF2-/- embryos or wild-type littermates, ER:H-RASV12 MEFs (from Julian Downward, London Research Institute, London, UK) and NIH 3T3 and HEK 293T (ATCC) cell lines were cultured in Dulbecco’s modified Eagle medium (DMEM, Life Technologies Inc.) supplemented with 10% fetal bovine serum (FBS) (HyClone). MEFs were transfected using Effectene (QIAGEN) and NIH 3T3 and HEK 293T cells using Polyfect (QIAGEN) according to the manufacturer’s instructions. Stable H, K, and N-RASD12- expressing NIH 3T3 cells were established by culturing transfected cells in 400 g/mL G418 (Sigma). Stable ER:H-RASV12-expressing MEFs and PP2A subunit-expressing HEK 293T cells were kind gifts from Julian Downward and Anne Claude Gingras (Samuel Lunenfeld Research Institute, Toronto, ON), respectively. Stable MEF, NIH 3T3-H-RASV12 and HEK 293T Arhgef2 knockdown cell lines were established by lentiviral infections of shRNA constructs. These viruses were produced by co-transfecting the HEK 293T packaging cell line with lentiviral shRNA hairpin plasmids targeting the murine or human ARHGEF2 gene and packaging plasmids pPAX2 and VSV-g using the CalPhos Mammalian Transfection Kit (Clontech). Lentiviral supernatants were collected, filtered and incubated with the target cells in the presence of 8g/ml Polybrene (Sigma). After 48h cells were subjected to puromycin (Sigma) selection (4g/ml for MEF and NIH 3T3-H-RASV12 cell lines and 2g/ml for HEK 293T cells) until all nontransduced cells died. All cultures were maintained in a 5% CO2 environment at 37oC.

Expression constructs: Full-length (Arhgef2 or Arhgef21-985), truncated (Arhgef287-151, Arhgef2236-572, Arhgef2236-433, Arhgef2473-572, Arhgef2473-985) and mutated (Arhgef2T243K) versions of murine and human ARHGEF2 cDNA (accession no. AF177032 and NM_004723.3, respectively) were subcloned into the pFlag-CMV2 (Sigma) or pEGFP-C1 (Invitrogen) vectors. Full-length murine p115RhoGEF cDNA (accession no. NM_001130150.1) was subcloned into pFlag-CMV2 vector. Murine ARHGEF2 pLKO.1 lentiviral shRNA constructs were obtained from The RNAi Consortium (TRC) and human ARHGEF2 shRNA sequences were cloned into the EcoRI and AgeI restriction sites of pLKO.1 (Table 1). A hairpin targeting GFP was used as a negative control (Table 1).

Table 2.1: Murine and Human Arhgef2 shRNA and GFP shRNA Sequences

Construct Forward sequence Reverse sequence mArhgef2 shRNA 1 5’- 5’- CCGGGCAGGAGATTTACAACCGAATCTCGA AATTCAAAAAGCAGGAGATTTACAACCGAATC GATTCGGTTGTAAATCTCCTGTTTTTG-3’ TCGAGATTCGGTTGTAAATCTCCTGTT-3’ mArhgef2 shRNA 2 5’- 5’- CCGGCCCTCATTTGTCCTACATGTACTCGAG AATTCAAAAACCCTCATTTGTCCTACATGTACT TACATGTAGGACAAATGAGGGTTTTTG-3’ CGAGTACATGTAGGACAAATGAGGGTT-3’ hArhgef2 shRNA 1 5’- 5’- CCGGAACCACGGAACTGGCATTACTCTCGA AATTCAAAAAAACCACGGAACTGGCATTACTC GAGTAATGCCAGTTCCGTGGTTTTTTTG-3’ TCGAGAGTAATGCCAGTTCCGTGGTT-3’ hArhgef2 shRNA 2 5’- 5’- CCGGAATGTGACTATCCACAACCGCCTCGA AATTCAAAAAAATGTGACTATCCACAACCGCC GGCGGTTGTGGATAGTCACATTTTTTTG-3’ TCGAGGCGGTTGTGGATAGTCACATT-3’ GFP shRNA 5’- 5’- CCGGTGCCCGACAACCACTACCTGACTCGA AATTCAAAAATGCCCGACAACCACTACCTGAC GTCAGGTAGTGGTTGTCGGGCA TTTTTG-3’ TCGAGTCAGGTAGTGGTTGTCGGGCA-3’

pCGT-H-, K-, N-RASV/D12, pCMV-Flag-AKAPLbc and pCMV-Flag-PP2A constructs were kind gifts from Dafna Bar Sagi (Langone Medical Centre, New York, NY), John Scott (Howard Hughes Medical Institute, Seattle, WA) and Anne-Claude Gingras, respectively. pCDNA3-Pyo- KSR-1 wild-type, mutant and truncated expression vectors were kind gifts from Deborah Morrison (Centre for Cancer Research, Frederick, MD) and were generated as described in Muller et al., 2001.

Cell treatments: ER:H-RASV12 MEFs were starved in DMEM containing 0% FBS for 16h and treated with 100nm 4-hydroxytamoxifen (4-OHT, Sigma) diluted in 100% ethanol. For MEK and PI3K inhibition experiments, MEFs and NIH 3T3-H-RASV12 cell lines were cultured in DMEM supplemented with 10% FBS and incubated with PD98059, UO126 or LY294002 (Sigma) diluted in DMSO (Sigma) for 48h. For immunofluorescence studies, MEFs were starved for 24h in DMEM containing 0% serum and treated in DMEM containing 10mM HEPES and 0.5mg/mL fatty acid-free bovine serum albumin (BSA) (A8806, Sigma). PDGF (Sigma) was suspended in Hank’s buffered saline solution (HBSS) containing 0.5mg/mL fatty acid-free BSA and 20 mM HEPES to a stock concentration of 1M.

Immunoprecipitations and Western blotting: For immunoprecipitation experiments, cells were scraped into ice-cold lysis buffer (30mM Tris pH7.5, 150mM NaCl, 1% Triton X-100, 0.2% sodium deoxycholate, 10mM NaF, 1mM Na3VO4 and 1mM PMSF) with Complete Protease Inhibitor cocktail (Roche) and cleared extracts incubated with protein-G sepharose and appropriate antibodies for 2h at 40C. Immunoprecipitates were washed three times with wash buffer (30mM Tris pH7.5, 300mM NaCl, 5mM NaF and 0.1% Triton X-100), resuspended in 2X sample buffer, boiled and protein complexes resolved by SDS-PAGE before transfer to PVDF (Imobilon) membranes for immunoblotting. For Western blotting, cells were scraped into ice- cold lysis buffer described above and incubated on ice for 20min, followed by centrifugation at 16,060xg at 4oC for 10min. Cleared lysates were resuspended in 2X sample buffer, boiled and proteins resolved by SDS-PAGE before transfer to PVDF membranes for immunoblotting.

RhoA and Rac1 activity assays: For pulldown experiments, active RhoA and Rac1 were assessed by incubation of cell lysates with GST-Rhotekin-RBD or GST-PAK-RBD, respectively (Cytoskeleton, CO, USA). Sub-confluent NIH 3T3-H-RASV12 cells stably expressing shGFP or shGEFm2 were serum-starved for 16h and lysed in ice cold HNMETG lysis buffer (50mM

HEPES pH 7.5, 150mM NaCl, 1.5mM MgCl2, 1mM EGTA, 1% Triton-X 100 and 10% glycerol). Lysates were clarified by centrifugation at 16,060xg at 4oC, equalized for total volume loading and rotated for 60min at 4oC with 20g of purified GST-RBD bound to glutathione Sepharose beads. The beads were washed three times with HNMETG wash buffer (50mM

HEPES pH 7.5, 300mM NaCl, 1.5mM MgCl2, 1mM EGTA, 0.1% Triton-X 100 and 10% glycerol) and processed for SDS-PAGE. For RhoA-GTP quantitation using RhoA G LISA kit

(Cytoskeleton, CO, USA), sub-confluent NIH 3T3-H-RASV12 cells stably expressing shGFP, shGEFm1 or shGEFm2 were serum-starved for 16h, washed, lysed in ice cold lysis buffer, cleared, snap-frozen in liquid nitrogen and stored at -70oC. Equal levels of total RhoA was confirmed with the Precision Red Advanced Protein Assay Reagent (Cytoskeleton) and lysates were processed for RhoA-GTP quantitation according to manufacturer’s protocol. Total GTP- bound RhoA was determined from cell lysates in triplicate and mean values from two independent experiments are shown +/- SD.

Antibodies: Polyclonal sheep anti-Arhgef2 murine antibodies were raised as described previously (Bakal et al., 2005). Monoclonal mouse anti-Arhgef human antibodies 3C5 and 14B11 were designed using N- and C-terminal human Arhgef2 peptides, respectively, and produced by hybridoma. Texas Red anti-mouse IgG (T-862) was obtained from Invitrogen. Western blotting and immunofluorescence were performed using the following primary antibodies: anti-RhoA (CST, 2117), anti-RAS (CST, 3965), anti-p44/42 MAPK (ERK1/2) (CST, 9102), anti-phospho- p44/42 MAPK (pERK1/2) Thr202/Tyr204 (CST, 9106), anti-MEK1/2 (CST, 9122), anti- phospho-MEK1/2 Ser217/221 (CST, 9154), anti-caspase 3 (CST, 9662), anti-cleaved caspase 3 (CST, 9661), anti-KSR-1 (gift from Deborah Morrison, see Cacace et al., 1999 for description of KSR-1 antibody generation), anti-phospho-KSR-1 S392 (CST, 2502), anti-PP2Ac (Millipore, 05-421), anti-RhoA (CST, 2117), anti-Rac1 (CST, 2465), anti-alpha tubulin (Molecular Probes), anti-Flag (M2, F3165, Sigma), anti-GFP (Invitrogen, G10362) and anti-Pyo (CST, 2448s). HRP- conjugated anti-mouse or anti-rabbit secondary antibodies were from GE Healthcare.

Quantitative PCR: RNA was extracted from NIH 3T3 or NIH 3T3-H-RASV12 cell lines using the RNeasy mini kit (QIAGEN). 100ng of RNA was converted into double-stranded cDNA at 42oC with SuperScript II RNase H-reverse transcription kit (Invitrogen). Quantitative PCR was performed with 50ng of template cDNA mixture from each cell line and murine Taqman gene expression assays for ARHGEF2 (Mm00434757_m1, Applied Biosystems) and TUBULIN (Mm00846967_g1, Applied Biosystems). Gene expression levels in the samples were calculated relative to control using the comparative CT method: CT = CTsample – CTcontrol, fold change = 2-CT. TUBULIN expression was used to normalize ARHGEF2 expression levels.

Luciferase reporter assays: The regulatory sequence of murine ARHGEF2 (nucleotides -62 to - 1968 upstream of the transcription start site (TSS)) was PCR-amplified from mouse BAC clones

33 and inserted into the pGL3 luciferase vector (pGL3pARHGEF2) (Promega, E1910). MEFs were co-transfected with pGL3pARHGEF2 and empty vector, pCGT-H-RASV12 or pCGT-K-RASD12 expression plasmids using LipoD293 (SignaGen, SL100668) and the luciferase activities were measured 24h after transfection using the Dual-Luciferase Reporter System (Promega) according to the manufacturer’s instructions.

Anchorage-independent growth: 60mm dishes were coated with bottom agar consisting of 0.6% ultra-pure agarose (Sigma), 2X DMEM, and 25% FBS and allowed to solidify at 40C for 30min. 1x105 cells were resuspended in top agar consisting of 0.4% agarose, 2X DMEM and 25% FBS 0 o at 37 C and poured over the bottom agar. After 24h at 37 C/5%CO2, 2ml of growth medium was 0 added to the top agar and was refreshed every 3 days. Cells were maintained at 37 C/5% CO2 for 10 days. For visualization, growth medium was removed and dishes were stained with 1ml of 0.0005% crystal violet in 70% ethanol for 4h at room temperature. Plates were washed with 70% ethanol and imaged at 10X or 40X on a dissecting microscope. Colonies greater than 2mm in diameter were counted manually at 10X magnification in triplicate. Results represent the mean of 3 independent experiments.

BrdU incorporation: NIH 3T3, NIH 3T3-H-RASV12, NIH 3T3-H-RASV12shGFP and NIH 3T3- H-RASV12shGEF1 and shGEF2 stable cell lines were plated at 1x103 cells/per well in a 96-well microplate in quadruplicate. BrdU reagent (Roche) was added to cells after 24h and incorporation was measured after 24h by colorimetric detection as per manufacturer’s protocol (Roche, 11647229001). Values reflect percentage BrdU incorporation relative to shGFP- expressing cells and represent the mean of three independent experiments.

(Olympus, Tokyo, Japan). Resolution was 512x512 with 12 bits/pixel. The following excitation wavelengths were used for enhanced GFP (473 nm) and Texas Red (559 nm). All images in each set of experiments were acquired with the same microscope sensitivity settings. All images compared within each figure panel were acquired on the same day, with identical staining conditions, gain and contrast setting, and same magnification. All statistical analyses were derived from 60 or more images from three independent experiments for each treatment condition.

Animal studies: All animal studies were carried out using protocols approved by the UHN Animal Care Committee. Xenograft studies in nude mice with NIH 3T3 cell lines were performed using 8-week old athymic NCr nude mice (Taconic Laboratories, Hudson, NY). Mice were allowed to acclimatize for one week in our institution’s animal care facility before being injected subcutaneously in the hip flank with 1 x 106 cells resuspended in 40ul of 1:1 PBS (Life Technologies) and growth factor-reduced matrigel (BD Biosciences). Mice were housed 3-4 to a cage and tumors were allowed to grow until they reached a maximum of 1.5cm in diameter or became ulcerated, at which point mice were sacrificed by carbon dioxide asphyxiation. Tumors were removed, weighed, measured, and fixed in OCT medium for histologic processing (described below). Five injections were performed per condition over four independent experiments. Tumor measurements were taken with a calliper and tumor volume was calculated by the ellipsoid formula V=/6 x (l x w2), where l and w denote the longest and shortest diameter, respectively.

Immunohistochemistry: For NIH 3T3 xenograft studies, tumor sections were fixed in Optimal Cutting Temperature (OCT) medium, flash frozen in methylbutanol, and stored at -80oC before being sent for immunohistological processing at Toronto General Hospital’s (TGH) Pathology Department. Tumor sections were probed for caspase 3 cleavage using anti-cleaved caspase 3 (Asp 175) antibody (CST 9661).

NMR-Based GEF assay: To quantify GEF activity in lysates of mammalian cells, nuclear magnetic resonance (NMR) was measured as described in Marshall et al., 2012 and Marshall et al., 2009. This assay monitors the heights of 1H-15N Heteronuclear Single Quantum Coherence (HSQC) peaks of 15N RhoA protein that are specific to either the GDP-bound or GTP-bound form. To measure nucleotide exchange, 2 mM GTPγS and 3.5 μl cleared lysate were added to a

35 μl sample of 0.2 mM 15N RhoA-GDP (residues 1–181) in NMR buffer (20 mM HEPES,

100 mM NaCl, 5 mM MgCl2, 2 mM Tris [2-carboxyethyl] phosphine [TCEP], 10% D2O [pH 7.0]). Nucleotide exchange was monitored by collecting successive 1H-15N HSQC spectra at 20°C using 4 or 8 scans (10 or 20 min/spectrum), depending on the reaction rate. Ten pairs of GDP/GTPγS-specific peaks (R5, V9, Q29, I46, A56, S73, Y74, D87, W158, T163) were used to evaluate the fraction of GDP-bound RhoA present at each time point, and the data were fitted to a single-phase exponential decay function to obtain the exchange rate, as described previously (Gasmi-Searbrook et al., 2010). To measure RhoA activity of truncated Flag-Arhgef287-151 and mutated Flag-Arhgef2T243K, plasmids containing these sequences were transfected into HEK 293T cells using Polyfect (QIAGEN) and NMR analysis was performed on lysates as described above.

Promoter analysis of ARHGEF2: Phylogenetic footprinting analysis was performed using mouse and human sequences for ARHGEF2 (NM_1162383.1 and NM_004723.3, respectively) (Zhang et al., 2003). Sequences were aligned to the genome with BLAT, where the TSS was ascertained and DNA 1kb downstream (3’) and 5kb upstream (5’) was pulled from the database. The 5kb and 1kb segments were analyzed separately using the Consite tool (Sandelin et al., 2004), employing all matrices found in the publicly-available Jaspar database. 3 and 1 cluster(s) of orthologous sequence areas were found in the 5kb and 1kb regions, respectively. In the 1kb region, sites were primarily linked to NFB transcription factor binding sites. When this region was expanded to include all Theria, 100% conservation was maintained. In the 5kb portion, the region just upstream of the TSS was linked to MYF, c-FOS and SAP-1 binding sites and were similarly well conserved across all Theria. H3K4me3 histone modifications were also analyzed by pull-down chip-seq data and showed a peak around the TSS in both mouse and human sequences. This peak overlaps with the putative NFB and MYF, c-FOS and SAP-1 binding sites present just downstream and upstream of the TSS, respectively, providing further evidence that these areas constitute the promoter region of ARHGEF2 (Ernst et al., 2011).

Statistical analyses: Values are expressed as means +/- standard deviation (SD) or +/- standard error (SE) as indicated. Paired Student’s t-tests (Kirkman, 2006) were performed to determine statistical significance between samples. Experiments were performed at least three times and means with p < 0.05 were considered statistically significant.

2.4 Results

2.4.1 Arhgef2 protein expression is acutely induced by the RAS/MAPK pathway

ARHGEF2 was identified as an upregulated gene in two independent studies looking at genome- wide transcriptional changes induced by oncogenic H-RAS and K-RAS in mouse fibroblast cells and human PDAC cells, respectively (Zuber et al., 2000, Qian et al., 2005). To discern whether Arhgef2 protein expression was increased in cells transformed by each RAS family member, we examined Arhgef2 levels in stable cell lines expressing H-RASV12, K-RASD12 and N-RASD12 compared to non-transformed isogenic fibroblasts (Figures 2.1A and 2.1B). Arhgef2 protein levels were upregulated in response to expression of each RAS family member and in proportion to RAS/MAPK pathway activation, as assessed by ERK1/2 phosphorylation (Figure 2.1B). We next determined whether Arhgef2 expression was a direct result of activated RAS or the secondary result of the transformed state. We used a murine embryonic fibroblast (MEF) cell line expressing a hydroxytamoxifen (4-OHT)-inducible form of H-RASV12 (ER:H-RASV12) (Gupta et al., 2007) that allowed us to examine Arhgef2 expression following the acute expression of H-RASV12. Arhgef2 expression increased within 2 hours of 4-OHT treatment compared to cells treated with vehicle control (Figure 2.1C). These data show that Arhgef2 is a direct target of H-RASV12.

To assess which RAS pathway regulates Arhgef2 expression, we treated RASV12-transformed fibroblasts with chemical inhibitors of the main branches of RAS signaling, the MAPK pathway and PI3K pathway. Treatment of H-RASV12-transformed mouse fibroblasts with the MEK inhibitors PD98059 (Figure 2.2A) or UO126 (Figure 2.2B) resulted in decreased Arhgef2 protein expression, whereas treatment with the PI3 kinase inhibitor LY294002 had no effect (Figure 2.2C), suggesting that Arhgef2 protein expression is regulated in response to MAPK pathway activation by oncogenic RAS.

Figure 2.1 Arhgef2 protein expression is acutely induced by oncogenic RAS. (A) Representative cell morphologies of mouse fibroblasts stably expressing empty vector or T7-H-RASV12, K-RASD12 or N-RASD12 family members. (B) Immunoblot analysis of Arhgef2 expression in mouse fibroblasts depicted in (A). RAS and phosphorylated ERK1/2 (pERK) levels represent RAS expression and pathway activation, respectively. Total ERK1/2 (ERK) and tubulin expression serve as protein loading controls. Data are representative of three independent experiments. (C) Immunoblot analysis of Arhgef2 expression following acute induction of H-RASV12. Mouse fibroblast cells stably expressing an estrogen receptor-tagged form of H-RASV12 (ER:H-RASV12) were serum starved for 16h followed by treatment with 100nM of 4-hydroxytamoxifen (4-OHT, upper panel) or vehicle control (EtOH, lower panel) over the indicated time periods. RAS induction and equal protein loading were confirmed by immunoblotting RAS and ERK1/2, respectively.

Figure 2.2 H-RASV12-induced Arhgef2 upregulation is dependent on MAPK pathway activation. (A) Mouse fibroblasts stably expressing T7-H-RASV12 were treated with DMSO or the MEK inhibitor PD98059 at the indicated concentrations for 48h. Changes in Arhgef2 expression were assessed by immunoblotting. Phosphorylated ERK1/2 represents the degree of RAS/MAPK pathway activation and total ERK1/2 and tubulin serve as protein loading controls. (B) H-RASV12-transformed fibroblast cells were treated with DMSO or the MEK inhibitor UO126 at the indicated concentrations for 48h and Arhgef2 protein expression was assessed by Western blot. Phosphorylated ERK1/2 levels represent the level of MEK inhibition and total ERK1/2 protein levels serve as gel loading controls. (C) H-RASV12-transformed fibroblast cells were treated with DMSO or the PI3K inhibitor LY294002 at the indicated concentrations for 48h and Arhgef2 protein expression was assessed by Western blot. Phosphorylated AKT (pAKT) denotes the level of PI3K inhibition and total AKT serves as a protein loading control.

2.4.2 ARHGEF2 is a transcriptional target of the RAS/MAPK pathway

To discern whether RASV12-mediated Arhgef2 upregulation occurred at the transcriptional level, we measured ARHGEF2 transcripts by quantitative PCR and found that they were elevated by two-fold in RASV12-transformed fibroblasts relative to wild-type cells (Figure 2.3A). To determine whether ARHGEF2 is a direct transcriptional target of RAS we identified a 1.9kb region upstream of the first exon of ARHGEF2 predicted to contain the putative promoter region, based on phylogenetic footprinting and CpG island enrichment, and cloned this region into a luciferase reporter (Figure 2.3B). Expression of H-RASV12 induced a 7-fold increase in ARHGEF2 promoter-mediated luciferase activity compared to cells expressing the ARHGEF2 promoter alone (Figure 2.3C, lanes 1 and 2). A similar level of the ARHGEF2 promoter activation was measured in response to K-RASD12 (Figure 2.3D, lanes 1 and 2). ARHGEF2 promoter activity was quenched with the MEK inhibitor PD98059 (Figure 2.3C), indicating that transcriptional activation of Arhgef2 requires MAPK pathway activation (Figure 2.3C, lanes 3

39 and 4). Together, these data show that Arhgef2 is a direct transcriptional target of the RAS/MAPK pathway.

Figure 2.3 ARHGEF2 is a transcriptional target of the RAS/MAPK pathway. (A) ARHGEF2 transcripts were quantified by real-time PCR in mouse fibroblasts stably expressing vector or T7-H-RASV12. Transcript levels were normalized to GAPDH and are represented as fold change over results from vector-only-expressing cells. Data represent the mean of three independent experiments +/- SD (p=0.00018). (B) Schematic representation of the putative promoter region of ARHGEF2. A 1907bp region upstream of the predicted transcription start site (TSS) was cloned into the pGL3 luciferase reporter for subsequent luciferase assays. (C) The ARHGEF2 luciferase reporter (pARHGEF2Luc) was co-expressed with empty vector or T7-H-RASV12 (lanes 1 and 2, respectively) and treated with the indicated concentrations of PD98059 for 16h (lanes 3 and 4, respectively). Luciferase activity was normalized to renilla expression and is represented as fold change over vector-expressing cells (upper graph). Data are representative of three independent experiments +/- SE. Cell lysates were assayed for RAS expression and MAPK pathway activity by immunoblotting for RAS and phosphorylated ERK1/2, respectively, and total ERK1/2 served as a protein loading control (lower panel). (D) Empty vector or T7-K-RASD12 expression plasmid was co- transfected with pARHGEF2Luc and harvested for luciferase reporter assays. Luciferase activity was normalized to renilla expression and is represented as fold change over results from vector-only-expressing cells (upper graph). Data are representative of three independent experiments +/- SE with p=0.00028. RAS expression and phosphorylated ERK1/2 were assessed by immunoblot as measures of RAS expression and RAS/MAPK activity, respectively, and total ERK1/2 served as a gel loading control (lower panel).

2.4.3 Arhgef2 is required for cell survival downstream of oncogenic RAS

To discern the functional significance of Arhgef2 in RAS-mediated cellular transformation, we stably knocked down Arhgef2 in murine fibroblasts transformed by RASV12 using two distinct ARHGEF2-directed lentiviral hairpins (Figure 2.4B, lanes 4 and 5). Depletion of Arhgef2 induced an apoptotic cell phenotype (Figure 2.4A), which was confirmed by immunoblotting for cleaved caspase-3 (Figure 2.4B). Moreover, Arhgef2 knockdown efficiency correlated with the degree of cell death in RASV12-transformed cells (Figure 2.4C).

In order to provide genetic support for the synthetic lethal interaction between Arhgef2 and RASV12, we examined the behaviour of RASV12 expression in murine embryonic fibroblasts (MEFs) derived from ARHGEF2-/- mice. Extensive cell death was observed in the Arhgef2-/- fibroblasts following RASV12 expression, whereas wild-type fibroblasts expressing RASV12 exhibited a refractile, transformed morphology with little change in cell viability (Figure 2.4D, columns 1 and 2). Moreover, re-expression of Arhgef2 in the ARHGEF2-/- fibroblasts expressing RASV12 restored cell survival, indicating that Arhgef2 is required for cell survival downstream of RASV12 (Figure 2.4D, column 3).

Figure 2.4 Arhgef2 is required for cell survival downstream of oncogenic RAS. (A) Representative cell morphologies of murine fibroblasts stably expressing H-RASV12 infected with a non-targeting hairpin (shGFP) or two distinct ARHGEF2 shRNAs (shGEF1 and shGEF2) and selected with puromycin for 48h. Arrowheads show rounded, pro-apoptotic cell morphologies of H-RASV12 transformed cells depleted of Arhgef2. (B) Cells described in (A) were lysed 5 days after infection and Arhgef2 depletion and caspase 3 cleavage were analysed by Western blotting with anti-Arhgef2 and anti-cleaved caspase 3 antibodies, respectively. Tubulin served as a protein loading control. (C) Murine fibroblasts stably expressing H-RASV12 were infected with shGFP, shGEF1 or shGEF2 and cell viability was determined by Alamar Blue staining after 72h (upper graph). Data are presented as percent viability compared to shGFP-expressing cells and represent the mean of four independent experiments +/- SE. ** denotes p<0.01 and * denotes p<0.05. 1000 cells were used per assay. Western blot analysis showing Arhgef2 expression in shGFP and shGEF-expressing cells quantified by Alamar Blue is shown (lower panel), with GAPDH serving as a protein loading control.

Expression of a cytoplasmically localized mutant of Arhgef2 exhibiting increased GEF exchange activity (Arhgef287-151, described in Meiri et al., 2012 and Figure 2.4E) with RASV12 rescued cell survival in an ARHGEF2-/- background (Figure 2.4D, column 4). However, expression of a catalytically inactive form of Arhgef2 (Arhgef2T247D, Meiri et al., 2012) also restored cell

42 viability in ARHGEF2-/- fibroblasts expressing RASV12, suggesting that the requirement for Arhgef2 for RASV12-mediated cell survival is independent of its enzymatic activity (Figure 2.4D, column 5). Furthermore, expression of p115RhoGEF (ARHGEF1), a close homologue to Arhgef2, and RASV12 in ARHGEF2-/- fibroblasts was unable to compensate for a loss of Arhgef2 expression, despite exhibiting high RhoA exchange activity by NMR analysis (Figure 2.4E). Together, these data demonstrate that RASV12 requires Arhgef2 expression, but not its RhoGEF activity, for cell survival.

Figure 2.4 Arhgef2 is required for cell survival downstream of oncogenic RAS. (D) Representative images of wild-type (ARHGEF2+/+, top row) and ARHGEF2 knockout (ARHGEF2-/-, bottom row) mouse fibroblasts transfected with free eGFP (vector, column 1), eGFP-H-RASV12 (column 2), eGFP-H-RASV12 and Flag-Arhgef2

(column 3), eGFP-H-RASV12 and Flag-Arhgef287-151 (column 4), eGFP-H-RASV12 and Flag-Arhgef2T247F (column 5) or eGFP-H-RASV12 and Flag-p115RhoGEF (column 6). Images were taken 4 days after transfection, following selection of plasmid-expressing cells with G418. (E) Real-time NMR measurement of RhoA nucleotide exchange in the presence of lysates from HEK 293T cells expressing eGFP, eGFP-Arhgef2, eGFP-Arhgef287-151, eGFP- Arhgef2E243K or eGFP-p115RhoGEF. As rate of nucleotide exchange for p115RhoGEF was 9.4-fold over Arhgef2 (r=0.132 vs r=0.014), its graphical representation is not to scale, as indicated by the breaks in the graph. Error bars represent +/- SD of a single experiment and data are representative of three independent experiments.

2.4.4 Arhgef2 contributes to RASV12-mediated cellular transformation in vitro and in vivo To determine whether Arhgef2 was required for RASV12-mediated cell transformation, we measured the ability of RASV12-transformed fibroblasts to support anchorage-independent growth in soft agar following knockdown of Arhgef2 (Figures 2.5A and 2.5B). Stable expression of RASV12 stimulated the growth of NIH 3T3 cells in soft agar with a mean number of 95 colonies/1000 cells whereas stable knockdown of Arhgef2 with one of two distinct shRNAs, reduced the number of colonies by 90% (n=9 colonies/1000 cells) compared to those cells expressing a non-targeting hairpin (82 colonies/1000 cells) (Figures 2.5A and 2.5C). To address the requirement of Arhgef2 in supporting tumor formation in RASV12-transformed fibroblasts, we generated subcutaneous xenografted tumors in NCr nude mice (Figure 2.5D). Parental and shGFP-expressing cells formed tumors reaching mean volumes of 600mm3 and 530mm3, respectively, within 10 days of injection (Figure 2.5D, left graph) whereas Arhgef2-depleted cells grew to a mean volume of 250mm3 and 200mm3 for shGEF1 and shGEF2, respectively. A two-fold-decrease in mean tumor weight was also measured in Arhgef2-depleted RASV12 xenografts compared to parental and hairpin controls (0.2g vs 0.5g, respectively) (Figure 2.5D, right graph). Moreover, Arhgef2-depleted tumors exhibited increased caspase 3 cleavage relative to parental and hairpin controls (Figure 2.5E). These data show that Arhgef2 is required for RASV12-mediated cell viability in vitro and in vivo.

The isolation of tumor cells derived from xenografts expressing Arhgef2 shRNAs revealed that in a subset of tumors, Arhgef2 expression was regained (Figure 2.6E, lanes 8 and 9 and Figure 2.6F, lane 8). We found that 3 of 9 tumors stably infected with Arhgef2 shRNA exhibited increased Arhgef2 protein levels compared to fibroblasts prior to injection. These data suggest that selective pressures may promote the acquisition of mutations or epigenetic changes at the

44 site of shRNA integration that result in promoter inactivation, resulting in the reconstitution of Arhgef2 expression. Alternately, contaminating macrophages or epithelial cells that contribute to tumor growth in murine xenografts may account for the presence of Arhgef2 protein expression.

Figure 2.5 Arhgef2 contributes to RASV12-mediated cellular transformation in vitro and in vivo. (A) Western blot analysis of Arhgef2 and RAS expression in murine fibroblast cell lines stably expressing empty vector, H-

RASV12, H-RASV12 and a hairpin control (shGFP, lane 3) or H-RASV12 and two distinct shRNAs targeting murine ARHGEF2 (shGEF1 and shGEF2, lanes 4 and 5 respectively). Total ERK1/2 served as a protein loading control. (B) Representative images of cell lines described in (A) resuspended in 0.3% agar to assess anchorage independent growth. (C) Mean colony number is depicted graphically and represents total number of colonies greater than 2mm in diameter per 60mm dish. Each experiment was performed in triplicate and results are the mean of three independent experiments +/- SE; Student’s t-test was used to generate p-values with p=7.6E-5 (shGEF1 vs shGFP) and p=1.0E-4 (shGEF2 vs shGFP) (** denotes p<0.01); 10000 cells were used per assay. (D) Representative images of NCr nude mice injected subcutaneously with 1x106 cells described in (A). Tumors were harvested when control tumors reached 1.5cm in diameter. Final tumor volumes and weights are depicted graphically and are the combination of four independent experiments and a total of n=21 tumors per condition. Error bars indicate +/- SE; Student’s t-test was used to generate p-values with p=0.0015 and p=0.0026 (shGFP vs shGEF1 and shGEF2 tumor volumes, respectively) and p=0.0078 and p=0.034 (shGFP vs shGEF1 and shGEF2 tumor weights, respectively) (**p<0.01, *p<0.05). (E) Representative images of immunohistochemical staining for cleaved caspase 3 in tumor sections derived from parental, shGFP, shGEF1 and shGEF2-expressing NIH 3T3-H-RASV12 xenografts. Images represent four tumors sampled from two independent experiments (n=8 per condition).

This highlights the requirement of Arhgef2 for RASV12-mediated tumor growth and suggests that tumors expressing mutant RAS positively select for high levels of Arhgef2 expression.

Figure 2.6 Arhgef2 protein expression is regained in a subset of Arhgef2-knockdown xenografts. (A, B) Western blot analysis of Arhgef2 expression in stable cell lines (described in Figure 2.5A) before injection into nude mice (lanes 1-5) and after harvesting from engrafted tumors (lanes 6-9). Each panel represents an independent experiment from n=4 experiments. Actin served as a protein loading control.

2.4.5 Arhgef2 contributes to the increased proliferative capacity of RASV12- transformed fibroblasts in a GEF-independent manner

Arhgef2 plays a role at several stages of cell-cycle progression in multiple cell types and is localized to the mitotic spindle of dividing cells (Aijaz et al., 2005, Bakal et al., 2005, Birkenfeld et al., 2007). Thus, we sought to determine whether Arhgef2 contributed to RASV12-mediated cell proliferation. Stable depletion of Arhgef2 in RASV12-transformed fibroblasts with both

46 hairpins resulted in a 20% decrease in cell proliferation as assessed by BrdU incorporation (Figure 2.7A). Expression analysis of cell cycle-associated genes in Arhgef2-depleted cells expressing RASV12 revealed that cyclin A expression was significantly reduced (Figure 2.7B). However, the expression of all other cyclins probed were unchanged, including the RhoA target cyclin D1. Moreover, expression of the cell cycle inhibitor p21/WAF1, known to be negatively regulated by RASV12-induced RhoA signaling (Sahai et al., 2002), was reduced in Arhgef2- depleted cells expressing RASV12 (Figure 2.7B, row 9). These results suggest that RhoA activity may not be significantly altered in RASV12-transformed fibroblasts depleted of Arhgef2 and that Arhgef2 may contribute to RASV12 function in a GEF-independent manner. This hypothesis was further supported by our earlier observations that a catalytically-inactive mutant of Arhgef2 was able to rescue RASV12-mediated cell survival in an Arhgef2-/- background (Figure 2.4D). To determine the effect of Arhgef2 depletion on Rho GTPase activity in RASV12-transformed cells, we probed for differences in the activities of RhoA and Rac1 in Arhgef2 knockdown cells compared to hairpin control-expressing cells using Rhotekin-Rho binding domain (Rhotekin- RBD) pulldown, RhoA G LISA and PAK-Rac binding domain (PAK-RBD) pulldown (Figures 2.7C-E). RhoA and Rac1-GTP levels were not significantly altered in Arhgef2 knockdown cells, demonstrating that changes in the downstream activation of Rho GTPases can not account for the requirement of Arhgef2 in RAS-mediated cellular transformation.

2.4.6 Arhgef2 is required for MAPK pathway activation in response to oncogenic RAS

To understand the mechanism underlying the contribution of Arhgef2 to RAS-mediated cellular transformation, we investigated whether elevated levels of Arhgef2 affected the signaling characteristics of upstream components of the RAS/MAPK pathway as part of a potential positive feedback mechanism. To that end, we expressed RASV12 in fibroblasts harboring stable knockdown of Arhgef2 and probed lysates for phosphorylated forms of MEK1/2 and ERK1/2 to assess MAPK pathway activity (Figure 2.8A). Both MEK1/2 and ERK1/2 were highly phosphorylated in RASV12-transformed fibroblasts expressing a non-targeting hairpin, while MEK1/2 and ERK1/2 phosphorylation was significantly reduced when Arhgef2 was depleted in these cells. A similar defect in RASV12-mediated ERK1/2 phosphorylation was seen in Arhgef2-/-

47 fibroblasts relative to wild-type fibroblasts (Figure 2.8B). Expression of an shRNA-resistant Arhgef2 cDNA (rArhgef2) or wild-type Arhgef2 restored MEK1/2 and ERK1/2 phosphorylation in response to RASV12 in Arhgef2 knockdown and Arhgef2-/- fibroblast cells, respectively, demonstrating that Arhgef2 is required for RASV12-induced activation of the MAPK pathway (Figure 2.8A, lane 7 and Figure 2.8B, lane 5).

Figure 2.7 Arhgef2 contributes to the proliferative capacity of RASV12-transformed fibroblasts in a GEF- independent manner. (A) NIH 3T3, NIH 3T3-H-RASV12 or NIH 3T3-H-RASV12 cell lines stably expressing shGFP, shGEF1 or shGEF2 were plated at 1000 cells/well in quadruplicate in 96-well plates and BrdU incorporation was measured over 24h. Results represent the percentage of BrdU incorporation relative to NIH 3T3-H-RASV12 cells and are the mean of three independent experiments +/- SE. (B) Western blot analysis of cell cycle-associated genes in uninfected H-RASV12-tansformed fibroblast cells (lane 1) or transformed cells stably infected with shGFP (lane 2), shGEF1 (lane 3) or shGEF2 (lane 4). Actin serves as a protein loading control. (C) Lysates derived from murine H-RASV12-transformed fibroblast cells stably expressing shGFP or shGEF2 were incubated with GST-tagged Rhotekin-Rho binding domain (RBD). Active RhoA-GTP in pulldowns (lanes 1 and 3) and total cellular RhoA (lanes 2 and 4) were detected by immunoblotting with anti-RhoA antibody. RhoA bound to RBD was normalized to total cellular RhoA for each condition (lower graph). Data are representative of three independent experiments. (D) Quantitation of RhoA-GTP levels in NIH 3T3-H-RASV12-transformed cells expressing shGFP, shGEF1 or shGEF2 by RhoA G LISA. Data are representative of two independent experiments +/- SD. (E) Lysates derived from murine H-RASV12-transformed fibroblast cells stably expressing shGFP, shGEF1 or shGEF2 were incubated with GST- tagged PAK-Rac binding domain. Active Rac1-GTP in pulldowns (first row) and total cellular Rac1 (second row) were detected by immunoblotting with anti-Rac1 antibody. Data are representative of two independent experiments.

To determine the specificity of Arhgef2-mediated MAPK pathway activation, we attempted to rescue the Arhgef2 knock down phenotype by expression of either AKAP-Lbc, the closest GEF family member to Arhgef2, or p115 RhoGEF, another RhoGEF family. Neither AKAP-Lbc (Figure 2.8A, lane 8) nor p115 RhoGEF (Figure 2.8B, lane 7 and Figure 2.4E) rescued MEK1/2 and ERK1/2 phosphorylation in response to acute RASV12 expression despite exhibiting high GEF activity, showing that Arhgef2 is uniquely required to mediate RAS-dependent activation of the MAPK pathway.

To determine whether Arhgef2-mediated MAPK pathway activation was dependent on its GEF activity, we co-expressed a catalytically inactive, shRNA-resistant form of Arhgef2

(rArhgef2T243K, Figure 2.4E) with RASV12 in fibroblasts depleted of endogenous Arhgef2 and found that MEK1/2 and ERK1/2 phosphorylation was fully restored (Figure 2.8A, lane 8). These findings were confirmed in Arhgef2-/- fibroblasts (Figure 2.8B, lane 6). These data show that Arhgef2 provides positive feedback loop for the RASV12/MAPK pathway in a manner independent of its GEF activity.

Figure 2.8 Arhgef2 is required for MAPK pathway activation in response to oncogenic RAS. (A) Mouse fibroblasts stably expressing shGFP, shGEF1 or shGEF2 were transfected with empty vector (lanes 1, 3 and 5) or H- RASV12 (lanes 2, 4 and 6) and assayed for ERK1/2 and MEK1/2 phosphorylation by Western blot. Rescue experiments were performed in shGEF2-expressing cells by co-transfecting H-RASV12 with Flag-rArhgef2 (shRNA resistant), Flag-Arhgef2E243K or Flag-AKAPLbc (lanes 7, 8 and 9, respectively). Expression of plasmids was confirmed by immunoblotting with anti-Arhgef2, anti-RAS and anti-Flag (AKAPLbc) antibodies and total levels of ERK1/2 and MEK1/2 served as protein loading controls. (B) ARHGEF2+/+ or ARHGEF2-/- MEFs were transfected with eGFP (lanes 1 and 3), eGFP-H-RASV12 (lanes 2 and 4) or co-transfected with eGFP-H-RASV12 and Flag- Arhgef2 (lane 5), eGFP-H-RASV12 and Flag-Arhgef2E243K (lane 6) or eGFP-H-RASV12 and Flag-p115RhoGEF (lane 7) and assayed for ERK1/2 activation by Western blot. Blots were probed with Arhgef2, Flag and RAS antibodies to confirm the expression from the transfected plasmids and total ERK1/2 and actin served as protein loading controls.

2.4.7 Arhgef2 is a component of the KSR-1 complex and is required for the dephosphorylation of its negative regulatory site on S392

Given that Arhgef2 catalytic activity is dispensable for RASV12-dependent MAPK pathway activation, we hypothesized that Arhgef2 may be providing a scaffold function for components of the MAPK pathway. First, we investigated whether Arhgef2 could form a complex with KSR- 1, a conserved MAPK scaffold that assembles pathway components into a large multiprotein complex required for efficient signal transduction. Analysis of Flag-Arhgef2 immune complexes from cells that expressed full-length or a series of Pyo-tagged KSR-1 deletions (Figure 2.9A) revealed that full-length KSR-1, KSR-1(1-539), KSR-1(1-424) and to a lesser extent KSR- 1(542-873), could interact with full-length Arhgef2 (Figure 2.9B, lanes 3, 4, 5 and 8). These data show that the C1 domain within the N-terminal half and the kinase domain of KSR-1 contribute to Arhgef2 binding. We next sought to determine whether the regulation of ERK1/2 activation by Arhgef2 depended on KSR-1 or signals to ERK1/2 through an alternative pathway. The microtubule unbound form of Arhgef2, Arhgef287-151, was expressed in wild-type or KSR1-/- fibroblasts (Figure 2.9C). Arhgef287-151 induced strong ERK1/2 phosphorylation in wild-type MEFs even in the absence of RASV12 expression (Figure 2.9C, lane 2), however, KSR-1-/- fibroblasts were resistant to Arhgef287-151-induced ERK1/2 phosphorylation (Figure 2.9C, lane 4). Co-expression of Arhgef287-151 and KSR-1, but not KSR-1 alone, restored ERK1/2 phosphorylation in KSR-1-/- cells (Figure 2.9C, lanes 5 and 6, respectively), demonstrating that Arhgef2 requires KSR-1 to positively regulate ERK1/2 activation.

Next, we investigated whether Arhgef2/KSR-1 binding affected the function of KSR-1. Growth factor or RASV12-induced KSR-1 function requires dephosphorylation of KSR-1 at 14-3-3 binding site S392 and subsequent translocation from the cytoplasm to the plasma membrane (Ory et al., 2003). We queried the requirement for Arhgef2 in growth factor-mediated KSR-1 translocation by stimulating wild-type or ARHGEF2-/- fibroblasts with PDGF and visualizing endogenous KSR-1 localization by immunofluorescence (Figure 2.10A). In 21.65% (21 of 97) of wild-type cells, KSR-1 translocated from the cytoplasm to the plasma membrane in a PDGF-

51 dependent manner (Figure 2.10A, columns 1 and 2 and Figure 2.10B). In contrast, in the absence of Arhgef2 only 3.45% of cells (3 of 87) underwent PDGF-dependent membrane translocation (Figure 2.10A, columns 3 and 4 and Figure 2.10B), a defect which was rescued by the expression of wild-type Arhgef2, with 29.59% of cells (29 of 98) showing KSR-1 plasma membrane localization (Figure 2.10A, columns 5 and 6 and Figure 2.10B).

Figure 2.9 Arhgef2 is a component of the KSR-1 complex and is required for the dephosphorylation of the negative regulatory site S392 on KSR-1. (A) Schematic representation of Pyo-tagged KSR-1 deletion constructs used to probe Arhgef2 binding in (B). (B) Pyo-tagged KSR-1 fragments depicted in (A) were co-expressed with Flag-Arhgef2 in HEK 293T cells. Complexes were immunoprecipitated with anti-Flag antibodies and proteins were detected by immunoblotting with anti-KSR-1 or anti-Flag (Arhgef2). (C) Immunoblot analysis of wild-type (WT) or KSR-1 deficient (KSR-1-/-) MEFs transfected with empty vector (lanes 1 and 3) or eGFP-Arhgef287-151 (lanes 2 and 4) and KSR-1-/- MEFs co-transfected with eGFP-Arhgef287-151 and Pyo-KSR-1 (lane 5) or Pyo-KSR-1 alone (lane 6). Lysates were assayed for activating phosphorylations of ERK1/2 by immunoblotting. Expression of endogenous and overexpressed proteins was determined by probing with anti-Arhgef2 and anti-KSR-1 antibodies and total ERK1/2 served as a protein loading control.

Since KSR-1 plasma membrane translocation requires dephosphorylation of S392 (Ory et al., 2003), we next determined if the non-phosphorylatable S392A point mutant form of KSR-1 could rescue the dependence on Arhgef2 for membrane translocation. We tested the capacity of wild-type or KSR-1S392A to translocate to the plasma membrane in ARHGEF2-/- fibroblasts. Whereas wild-type KSR-1 was largely unable to translocate to the plasma membrane (8.70% or 6 of 69 cells, Figure 2.10C and Figure 2.10D), KSR-1S392A underwent greatly increased plasma membrane localization even in the absence of Arhgef2 (36.84% or 28 of 76 cells) (Figure 2.10C, columns 1 and 2 and Figure 2.10D). These data show that Arhgef2 is required for translocation of KSR-1 to the plasma membrane in a manner that depends on the dephosphorylation of KSR- 1S392. Lastly, we showed that re-expression of Arhgef2 and KSR-1 in Arhgef2-/- fibroblasts was insufficient to induce membrane translocation of KSR-1 in the absence of PDGF treatment (5.97% or 4 of 67 cells) (Figure 2.10C, column 3 and Figure 2.10D). Moreover, the requirement for growth factor stimulated KSR-1 translocation to the plasma membrane could be subverted by the expression of Arhgef2∆87-151, with 29.58% of cells (21 of 71) exhibiting KSR-1 plasma membrane localization (Figure 2.10C, column 4 and Figure 2.10D) (Meiri et al., 2012). Both Arhgef2∆87-151 and KSR-1 localized to the plasma membrane in the absence of PDGF stimulation (Figure 2.10C, column 4, lower and upper panels, respectively). These data suggest that the growth factor dependence of KSR-1 translocation may be conferred by the release of Arhgef2 from the microtubule array. To determine whether Arhgef2 regulation of the RASV12/MAPK cascade is coupled to the dephosphorylation of KSR-1, we asked whether wild-type KSR-1 or KSR-1S392A could restore RASV12-induced ERK1/2 phosphorylation in the absence of Arhgef2. We expressed shRNA- resistant Arhgef287-151, (rArhgef287-151) together with either wild-type KSR-1 or KSR-1S392A in Arhgef2 knockdown fibroblasts (Figure 2.10E). As previously shown, RASV12 expression induced ERK1/2 phosphorylation in hairpin control-expressing fibroblasts but not fibroblasts depleted of Arhgef2 (Figure 2.10E, lanes 2 and 4). High expression of rArhgef2 in Arhgef2- depleted cells greatly enhanced ERK1/2 activation in response to RASV12, supporting the model that increased levels of Arhgef2 results in amplification of the ERK1/2 cascade (Figure 2.10E, lane 5). Importantly, expression of KSR-1S392A was able to restore RASV12-mediated ERK1/2 phosphorylation in Arhgef2 knockdown cells (Figure 2.10E, lane 6). However, wild-type KSR-1 was unable to fully restore ERK1/2 activation to the same extent as KSR-1S392A despite similar

Figure 2.10 Arhgef2 is required for plasma membrane translocation of KSR-1. (A) Immunofluorescence analysis of endogenous KSR-1 localization and eGFP or eGFP-Arhgef2 expression in ARHGEF2+/+ (columns 1 and

2) or ARHGEF2-/- MEFs (columns 3-6) transfected with free eGFP (columns 2 and 4) or eGFP-Arhgef2 (column 6) and treated with vehicle control (BSA, columns 1 and 3) or 25ng/ml PDGF for 10 min (columns 2, 4 and 6). Arrows indicate plasma membrane localization of KSR-1 (columns 2 and 6, top row) and eGFP-Arhgef2 (column 6, bottom row) in the transfected cells. Images are representative of four independent experiments. (B) Quantification of the number of cells exhibiting KSR-1 membrane translocation in response to BSA, PDGF and/or eGFP-Arhgef2 in ARHGEF2+/+ or ARHGEF2-/- MEFs. Data are represented as percentage of cells analyzed in each condition and are the combination of three independent experiments. (C) Immunofluorescence analysis of ARHGEF2-/- MEFs co- transfected with wild-type KSR-1 (columns 1, 3 and 4) or KSR-1S392A (column 2) and free eGFP (columns 1 and 2), eGFP-Arhgef2 (column 3) or eGFP-Arhgef287-151 (column 4). Cells were fixed and stained for KSR-1 (top row) or eGFP (bottom row). Arrows indicate plasma membrane localization of KSR-1 (columns 2 and 4) and eGFP-Arhgef2 (column 4). Images are representative of four independent experiments. (D) Quantification of the number of cells exhibiting KSR-1 membrane translocation in response to pyo-KSR-1, pyo-KSR-1S392A, pyo-KSR-1 + eGFP-Arhgef2 or pyo-KSR-1 + eGFPArhgef287-151 in ARHGEF2-/- MEFs. Data are represented as percentage of cells analyzed in each condition and are the combination of three independent experiments. (E) Western blot analysis of murine fibroblasts stably expressing shGFP, shGEF1 or shGEF2 transfected with empty vector (lanes 1 and 3) or H-RASV12 (lanes 2 and 4) or co-transfected with H-RASV12 and Flag-rArhgef2 (lane 5), Pyo-KSR-1S392A (lane 6) and wild-type Pyo-KSR-1 (lane 7). Inhibitory phosphorylation of KSR-1 on S392 was assessed using a phospho-KSR-1S392- specific antibody (KSR-1pS392) and ERK1/2 activation was detected with anti-phospho-ERK1/2. Anti-Arhgef2, KSR-1 and RAS antibodies detected the expression of transfected plasmids and total ERK1/2 served as a loading control. expression levels (Figure 2.10E, lane 7). These data show that dephosphorylation of S392 of KSR-1 is sufficient to overcome the Arhgef2 dependence of RASV12-mediated ERK1/2 activation in fibroblasts.

2.4.8 Arhgef2 is required for PP2A-mediated dephosphorylation of KSR-1 on S392

We have identified Arhgef2 as a PP2A interacting partner in a proteomic screen designed to probe for proteins that bound to the PP2A catalytic subunit (Meiri et al., manuscript in submission) and we found that Arhgef2 interacts with the B’ regulatory PP2A subunits (PPP2R5A, PPP2R5B and PPP2R5E). To determine the mechanism underlying the dependence of KSR-1S392 dephosphorylation on Arhgef2, we hypothesized that Arhgef2 may act as a bridge between KSR-1 and PP2A, since S392 dephosphorylation by PP2A in response to growth factor stimulation has previously been described (Ory et al., 2003).

First, we confirmed the previously published data showing an interaction between KSR-1 with the B’ regulatory PP2A subunits (Figure 2.11A) (Ory et al., 2003). We observed that Arhgef2

55 bound the same PP2A subunits that interact with KSR-1 (Figure 2.11A). We evaluated the regions of Arhgef2 involved in PP2A and KSR-1 binding by expressing deletion mutants of Arhgef2 and probing for the catalytic subunit of PP2A and KSR-1 in Arhgef2 immune complexes (Figure 2.11B). Analysis of Arhgef2 immunoprecipitates revealed that endogenous KSR-1 interacted with full-length Arhgef2, Arhgef2(236-572) and Arhgef2(236-433). These results localize the binding site for KSR-1 to the DH domain of Arhgef2 (Figure 2.11C), while PP2Ac binds to the Arhgef2 PH domain (Figure 2.11C). These data show that KSR-1 and PP2A bind to distinct sites on Arhgef2 and suggest a model by which Arhgef2 may link KSR-1 to PP2A.

To determine whether Arhgef2 is a scaffold that links KSR-1 to PP2A, we assessed the requirement of Arhgef2 for the KSR-1/PP2A interaction. To that end, we stably infected PP2A B subunit-expressing cells with Arhgef2 shRNA and probed PP2A immune complexes for KSR-1 (Figure 2.11D). Knockdown of Arhgef2 was confirmed by immunoblotting total cell lysates (Figure 2.11D, fourth row). KSR-1 was detected in PPP2R5A, PPP2R5B and PPP2R5E (B’ subunit), but not PPP2R2A (B subunit) immune complexes. However, in Arhgef2-depleted cells, KSR-1 could not be detected in any of the PP2A B’ subunit complexes. These data show that the interaction between KSR-1 and PP2A is dependent on Arhgef2, providing a model whereby Arhgef2 serves as a scaffold to recruit the PP2A B’ subunits required for the dephosphorylation of the negative regulatory S392 site on KSR-1 and activation of the MAPK pathway.

Figure 2.11 Arhgef2 is required for PP2A-mediated dephosphorylation of KSR-1 on S392. (A) Flag-tagged PP2A catalytic (lane 1) and regulatory (lanes 2-5) subunits were stably expressed in HEK 293T cells and PP2A immune complexes were isolated using anti-Flag antibodies (row 1). PP2A complexes were probed for endogenous Arhgef2 and KSR-1 (rows 2 and 3). Expression levels of Arhgef2 and KSR-1 in whole cell lysates are indicated in rows 4 and 5. (B) Schematic representation of Arhgef2 constructs used in (C). (C) Flag-tagged Arhgef2 fragments were expressed in HEK 293T cells and complexes were immunoprecipitated with anti-Flag antibody. Complexed proteins were detected by immunoblotting with anti-KSR-1 or anti-PP2Ac antibodies and whole cell lysates were probed for KSR-1 and PP2Ac expression. (D) HEK 293T cells stably expressing Flag-tagged PP2A regulatory subunits PPP2R5A, PPP2R5B, PPP2R5E and PPP2R2A were infected with a hairpin control (shGFP) or shRNA against human ARHGEF2 (shGEF). PP2A subunits were immunopurified with anti-Flag (row 1) and probed for the presence of endogenous KSR-1 (row 2). Arhgef2 knockdown and PP2A subunit expression was confirmed by immunoblotting whole cell lysates with anti-Arhgef2 and anti-Flag, respectively.

2.5 Discussion

In this chapter, we have uncovered a positive feedback loop in which MAPK-dependent increases in Arhgef2 expression potentiate the MAPK cascade in RASV12-transformed cells (Figure 2.12). We found that Arhgef2 is transcriptionally upregulated by the RAS/MAPK pathway and in turn positively regulates MAPK activation by facilitating the PP2A-mediated dephosphorylation and activation of the MAPK scaffold, KSR-1, in response to RASV12. Arhgef2 functions independently of its GEF activity and acts as an adaptor molecule between the PP2A B’ subunit family and KSR-1. Ablation of Arhgef2 prevents RASV12- and KSR-1- mediated MEK1/2 and ERK1/2 activation, induces apoptosis and reduces the growth of RASV12- induced xenografts. Together, these data provide mechanistic insight into the regulation of MAPK signaling downstream of oncogenic RAS and propose a novel mechanism by which RAS mutated cancers may select for increased MAPK survival signaling.

Figure 2.12 The Arhgef2/PP2A complex provides a positive feedback loop to the KSR/MAPK pathway in RASV12-transformed cells. Schematic model showing the induction of ARHGEF2 transcripts in response to MAPK activation by oncogenic RAS. In the presence of RASV12 Arhgef2 can recruit the B’ subunit of PP2A to the inactive

KSR-1 complex, thereby facilitating its dephosphorylation on S392. KSR-1 can then translocate to the plasma V12 membrane, bringing Raf/MEK/ERK into close proximity and enhancing RAS -induced MAPK signaling.

The precise mechanism of ARHGEF2 transcriptional regulation downstream of the RAS/MAPK pathway remains to be elucidated. One possibility is modulation via the transcription factor hPTTG1, previously shown to regulate ARHGEF2 in breast cancer, as several studies have indicated that activation or inhibition of the MAPK pathway can induce or repress hPTTG1 expression, respectively (Liao et al., 2012, Vlotides, 2006, Hernandez, 2008). Since the putative promoter region of ARHGEF2 used in this study contains the hPTTG1 binding region, it is possible that RAS/MAPK activation upregulates ARHGEF2 through an indirect mechanism involving hPTTG1. Analysis of the promoter region of ARHGEF2 by phylogenetic footprinting, however, revealed three conserved clusters of alternative transcription factor binding sites (described in Experimental Procedures). One region lies immediately in front of, and another downstream of, the transcription start site and contains putative myf, c-fos and SAP-1 and NFB binding sites, respectively. c-fos, SAP-1 and NFB are activated by the RAS/MAPK pathway and are therefore potential mediators of RASV12-induced ARHGEF2 upregulation (Wang et al., 2000, Galanis et al., 2001, Schulze-Osthoff et al., 1997).

Arhgef2 has been implicated in cell survival in several contexts. Arhgef2 is activated by TNF in tubular epithelial cells and was shown to mediate cell survival in response to TNF, hyperosmotic shock, taxol and EGF by inducing the post-translational stabilization of p21 (Kakiashvili et al., 2011, Nie et al., 2012). Lung cancer cells harboring mutations in p53 exhibited dose-dependent decreases in cell viability in response to Arhgef2 depletion (Mizuarai et al., 2006). In the aforementioned studies, Arhgef2 was shown to affect cell survival and/or proliferation via its GEF activity toward RhoA. Although RASV12 activates RhoA, we did not detect a significant change in RhoA activity upon stable depletion of Arhgef2 in RASV12- expressing fibroblasts (Qiu et al., 1995). These data show that oncogenic RAS induces RhoA- GTP independently of Arhgef2 and that Arhgef2 is likely not contigent on RASV12-mediated survival signaling by altering Rho GTPase levels. This is in agreement with Chen et al., who found that increased RhoA-GTP levels induced by RASV12 were due to a decrease in p190RhoGAP activity, with little change in total cellular Rho GEF activity or Rho-Rho GDI immune complexes (Chen et al., 2003). Instead, oncogenic RAS promotes the adaptor function of Arhgef2, perhaps by inducing its re-localization from RhoA to PP2A B’ subunit and KSR-1

59 pools in the cell through yet unidentified mechanisms. One interesting possibility is that oncogenic RAS perturbs microtubule dynamics, resulting in the release of Arhgef2 from the microtubule array. This hypothesis is supported by our observations that co-expression of KSR-1 and an Arhgef2 mutant incapable of binding microtubules (Arhgef287-151) can potentiate MAPK activation in the absence of growth factor stimulation or oncogenic RAS. Moreover, these data implicate Arhgef2 in the interaction between anti-mitotic chemotherapeutic drugs (AMCDs) and the inhibition of RAS/MAPK signaling. These agents are widely used for the treatment of solid malignancies and act by interfering with microtubule dynamics, resulting in cell cycle arrest and apoptosis (Jordan and Wilson, 2004, Wilson et al., 1999). MAPK signaling has been shown to be modulated by AMCDs and is implicated in AMCD resistance (Shinoharah-Gotoh et al., 1991, Orr et al., 2005). Thus, one might speculate that AMCD-induced Arhgef2 release from microtubules may contribute to the acquired resistance of cancer cells harboring RAS mutations. Alternately, Arhgef2 may contribute to inhibition of MAPK signaling in AMCD-responsive tumor cells. This is highly dependent on whether the AMCD in question induces the sequestration or release of Arhgef2 from the microtubule array, as different classes of AMCDs can act by destabilizing or stabilizing microtubules and would thereby impinge on Arhgef2 regulation in opposing manners. Ascertaining the relationship between oncogenic RAS, microtubule regulation and different AMCDs would therefore be an interesting area of future study.

Although we cannot exclude the possible contribution of subtle Arhgef2-mediated changes in RhoA activity to increased cell survival downstream of oncogenic RAS, Arhgef2-dependent KSR-1 regulation can account for the discrepancy between a mild decrease in RhoA activity and strong inhibition of cell survival and transformation in Arhgef2-depleted cells. KSR-1 is critical for cell survival in EGFR and oncogenic RAS-dependent tumors via activation of the MAPK pathway (Xiao et al., 2010). Furthermore, KSR-1 is required for cellular transformation in response to oncogenic RAS and this is strictly dependent on the dephosphorylation of KSR-1 at S392 by PP2A (Kortum et al., 2004, Joneson et al., 1998, Razidlo et al., 2004, Nguyen et al., 2002, Ory et al., 2003). KSR-1 has also been shown to mediate TNF-induced cell survival in intestinal epithelial cells through the activation of ERK1/2, suggesting that Arhgef2 and KSR-1 may cooperate in other cellular contexts (Yan et al., 2001, Yan et al., 2004).

Until now, the mechanism of growth-factor induced recruitment of the regulatory subunit of PP2A to KSR-1 in mammalian cells was unknown (Ory et al., 2003). Here, we provide evidence that Arhgef2 functions as an adaptor protein, linking the PP2A B’/PR61/B56/R5 subunit to the KSR-1/PP2A complex in response to RAS activation, thereby potentiating oncogenic signaling. The functional significance of these interactions is supported by genetic evidence in C. elegans, where the regulatory subunit of PP2A was found to be a critical activator of PP2A-mediated RAS signaling (Sieburth et al., 1999, Kao et al., 2003). These data therefore place Arhgef2 at a critical point in the regulation of KSR-1. Although PP2A has been shown to negatively regulate MAPK signaling at the level of MEK1/2 and ERK1/2, it is unlikely that Arhgef2 mediates these interactions since it has been shown to require the B/PR55/R2 family of regulatory PP2A subunits and not the B’/PR61/B56/R5 subunit family shown to bind Arhgef2 and KSR-1 in this study (Zhou et al., 2002, Sontag et al., 1993, Silverstein et al., 2002 and Meiri et al., in submission). In support of this observation, only depletion of the B subunit of PP2A was shown to activate ERK1/2 signaling in Drosophila Schneider 2 cells, while depletion of the B’ alpha and beta subunits induced apoptosis (Silverstein et al., 2002). These data further substantiate a role for Arhgef2-mediated KSR-1 function in cell survival downstream of RASV12 and suggest that by initiating the formation of PP2A/B’ subunit complexes, increased Arhgef2 expression may favor the positive regulation of the MAPK cascade in RASV12-transformed cells.

The recruitment of PP2A to KSR-1 by Arhgef2 may also explain the requirements of the PH and DH domain of Arhgef2 for cellular transformation (Whitehead et al., 1995). It was originally speculated that the PH domain is responsible for targeting Arhgef2 to the plasma membrane, where it could exert its exchange activity on RhoA; however, subsequent studies have not substantiated a requirement for plasma membrane localization of Arhgef2 for its catalytic activity. Considering that the PH domain and DH domains of Arhgef2 are required for its interaction with PP2A and KSR-1, respectively, one may speculate that Arhgef2 partially elicits its classical transforming ability via the activation of KSR-1. This is in agreement with the observation that while wild-type KSR-1 is unable to transform fibroblasts independently, moderate overexpression of a double S392/T274 mutant of KSR-1 can induce anchorage- independent growth in the absence of RASV12 (Razidlo et al., 2004).

The results presented in this chapter unveil an essential role for Arhgef2 in RASV12-mediated cellular transformation in fibroblast cells. A critical question that remains, however, is whether the function of Arhgef2 in RASV12-induced fibroblast transformation is paralleled in human epithelial tumors harboring endogenous RAS mutations. Moreover, it is of considerable interest to determine if Arhgef2 contributes to the malignant conversion of RAS-mutated tumors, where current therapies are most ineffective. To begin to answer these questions, in the next chapter I will query the role of Arhgef2 in human epithelial models of RAS tumorigenesis and its effect on their epithelial-to-mesenchymal (EMT) conversion.

Chapter 3

Arhgef2 is Required for Primary Tumorigenesis and Promotes Mesenchymal Transition in Pancreatic Ductal Adenocarcinoma

3.1 Abstract

Mutations in K-RAS are present in 20% of human solid malignancies, including 95% of pancreatic ductal adenocarcinomas (PDACs), 50% of colorectal tumors and 30% of non-small- cell lung cancers (NSCLCs) (Maitra et al., 2003, Johnson et al., 1993, Rodrigues et al., 1990). PDAC has one of the shortest five-year survival rates, exhibiting overwhelming resistance to currently available therapies (Coz et al., 2002). In this chapter, we find that Arhgef2 is required for proliferation and survival across several RAS-mutated human epithelial cancer cell lines. Arhgef2 contributes to primary tumorigenesis in PDAC xenograft models and potentiates MAPK signaling in these cells by a parallel mechanism to that observed in fibroblasts. Furthermore, analysis of human tumor microarrays (TMAs) revealed that Arhgef2 protein expression correlates with progressive tumor grade in PDAC, colorectal and NSCLC cancers, implicating Arhgef2 in the malignant conversion of RAS-mutated tumors. Indeed, we find that Arhgef2 promotes a mesenchymal morphology in PDAC and NSCLCs harboring RAS mutations. Arhgef2 depletion results in a reversion to an epithelioid gene signature, cell morphology and increased E-cadherin protein expression. Moreover, Arhgef2 is required for epithelial-to- mesenchymal transition (EMT) in a murine mammary epithelial cell model of TGF-induced EMT. Together, these data implicate Arhgef2 at multiple stages of RAS tumorigenesis and suggest that Arhgef2 may be an effective therapeutic target in PDAC and other cancers.

3.2 Introduction

Over 85% of human cancers arise from epithelial cells, making them pertinent model systems to study the mechanisms contributing to human tumorigenesis. Epithelial cells are cuboidal, polarized cells that associate closely together to form an organized, compact epithelium that lines the cavities and surfaces of structures throughout the body. The epithelium is held together through several types of interactions, including zona occludens-1 (ZO-1) containing tight junctions, E-cadherin-based adherens junctions, desmosomes and gap junctions (Radisky, 2005). Together, these epithelial sheets form protective barriers against external environmental hazards and foster physiologically defined subdomains within different organs of the body.

Epithelial cancers, or carcinomas, progress in a multistep fashion via the sequential accumulation of genetic lesions within an epithelial cell (Figure 3.1) (Weinberg, RA, 1989). According to this paradigm, each oncogenic event confers the tumor cell with transforming properties that culminate in a fully malignant tumor. In colorectal and pancreatic cancer, this step-wise genetic model has been phenotypically aligned with graded yet distinct neoplastic changes in the tissue with progressive stages of tumorigenesis (Fearon et al., 1990, Hruban et al., 2001). The consecutive pre-invasive stages of pancreatic tumorigenesis have been morphologically categorized as pancreatic intraepithelial neoplasias 1A, 1B, 2, 3 (PanIN-1A, -1B, PanIN-2, PanIN-3) and advanced pancreatic ductal adenocarcinoma (PDAC), and are genetically defined by the sequential gain or loss-of-function mutations of K-RAS, p16/INK4, p53 and smad4, respectively (Hruban et al., 2001, Schneider et al., 2003, Apple et al., 1999, Wilentz et al., 1998, DiGiuseppe et al., 1994, Wilentz et al., 2000). K-RAS mutations are considered the primary initiating event in PDAC, as they are commonly found in pre-neoplastic tissues (Klimstra et al., 1994, Tada et al., 1996). p53 overexpression and smad4 loss, however, are detected late in PDAC progression and drive its malignant conversion (DiGiuseppe et al., 1994, Wilentz et al., 1998).

Epithelial-to-mesenchymal-transition (EMT) is a defining feature of late-stage tumorigenesis, as it enables the invasion of tumor cells through the basement membrane and to distant organs of the body. EMT is characterized by the dissolution of epithelial cell junctions, a loss of cell

Figure 3.1 Multistep tumorigenesis in pancreatic ductal adenocarcinoma. Progressive changes in pancreatic ductal epithelial cell growth and architecture are demonstrated schematically (above) and immunohistologically (below, with epithelial cell layer stained with keratin in dark blue). PanIN-1A predominantly develops following mutations in K-RAS, with subsequent losses in p21, p16, p53 and SMAD4 tumor suppressors contributing to neoplastic progression (PanIN-1B-PanIN-3) and ultimately, malignant conversion (ADC) (adapted from Ma et al., 2011). adhesion and polarity, and the acquisition of a motile, mesenchymal phenotype (Guarino et al., 2007). At the molecular level, mesenchymal conversion involves the downregulation or delocalization of junctional proteins such as E-cadherin and ZO-1 and the upregulation of mesenchyme-defining genes like vimentin (Peinado et al., 2004). E-cadherin loss is the most prominent feature of EMT and is sufficient to induce the full mesenchymal transition of an epithelial cell by decreasing cell-cell adhesion and promoting invasion and motility (Lehembre et al., 2008, Thompson et al., 1994). Mesenchyme-specific transcription factors are also induced, resulting in widespread gene expression changes to sustain the mesenchymal phenotype (Taube et al., 2010, Kim et al., 2010, Hoshida et al., 2009).

Transforming growth factor- (TGF) regulates a diverse number of processes including cell proliferation, differentiation, apoptosis, and EMT (Shi et al., 2003). TGF ligands function by binding type I TGF serine/threonine kinase receptors, triggering their dimerization with type II receptors, and the subsequent phosphorylation and activation of smad proteins. Membrane- associated smad7 activates the cytoplasmic smads 2 and 3, which then cooperatively activate

65 smad4, resulting in its nuclear translocation and the transcription of target genes (Shi et al., 2003). TGF has been shown to drive EMT via smad-dependent and independent pathways, the latter of which include RAS, JNK, p38, ERK, PI3K and RhoA signaling (Oft et al., 1996, Atfi et al., 1997, Hartsough et al., 1995, Bakin et al., 2000, Bhowmick et al., 2001). TGF can drive mesenchymal transition through the reorganization of cytoskeletal components and breakdown of cell-ECM interactions and through the activation of EMT-inducing transcription factors such as NFB, snail, and slug (Janda et al., 2002, Ozdamar et al., 2005, Huber et al., 2004, Peinado et al., 2003, Peinado et al., 2004).

The Rho GTPases have been linked to EMT via TGF-dependent and independent mechanisms. Rho proteins positively regulate motility, migration and invasion by modulating the actin cytoskeleton and by regulating cell adhesion in fibroblast cells (Ridley et al., 1992, Kaibuchi et al., 1999). In addition, RhoA regulates the formation of cadherin-based cell-cell contacts in epithelial cells (Braga et al., 1999). RhoA activation has been established in the reversion of an epitheloid phenotype toward a migratory, fibroblastoid phenotype in NIH 3T3 cells and is required for TGF-induced mesenchymal transition in a mammary epithelial cell model of EMT (Sander et al., 1999, Bhowmick et al., 2001). RhoA promotes the invasive phenotypes of tumor cells and contributes to metastases in xenograft tumor models, highlighting its essential role in tumor progression (Yoshioka et al., 1998, Yoshioka et al., 1999).

Arhgef2 activates RhoA and has been shown to localize to and regulate the permeability of epithelial and endothelial tight junctions (Benais-Pont et al., 2003, Birukova et al., 2006, Guillemot et al., 2008). Increased expression of Arhgef2 at the apical junctions of epithelial cells results in junctional disassembly, thereby compromising epithelial barrier function (Samarin et al., 2007). In fibroblast cells, Arhgef2 initiates stress fiber and focal adhesion formation via its exchange activity on RhoA, thereby regulating migration and cell adhesion (Krendel et al., 2002, Callow et al., 2005, Nalbant et al., 2009, Guilluy et al., 2011). Furthermore, Arhgef2 is a transcriptional target of TGF and mediates TGF-induced migration in retinal epithelial cells in a RhoA-dependent manner (Tsapara et al., 2010). Arhgef2 has also been shown to promote the metastatic potential of breast cancer cells overexpressing hPTTG1 by increasing their invasive properties (Liao et al., 2012).

In chapter 2, we identified ARHGEF2 as a transcriptional target of oncogenic RAS in fibroblast cells that was essential for RAS-mediated survival and transformation. Given the high proportion of RAS mutations in PDAC, we hypothesized that Arhgef2 levels may be elevated in pancreatic tumors and contribute to primary tumorigenesis. Moreover, we predicted that Arhgef2 may contribute to EMT by promoting the dissolution of cell junctions and increasing their migratory properties.

3.3 Experimental Procedures

Cell lines and cell culture: HEK 293T, PANC-1, HPAF-II, PL-45, (from ATCC), H1264, DLD1, DK04 (from Ming Sound-Tsao, Princess Margaret Hospital, Toronto, ON) and NMuMG (from Jeff Wrana, Samuel Lunenfeld Research Institute, Toronto, ON) cell lines were cultured in Dulbecco’s modified Eagle medium (DMEM, Life Technologies Inc.) supplemented with 10% fetal bovine serum (FBS) (HyClone). SK-OV-3 (from Gordon Mills, The University of Texas, MD Andersen Centre, Houston, Texas), BxPC3, H727, A549 (ATCC) and H520 (from Ming Sound-Tsao) cell lines were cultured in RPMI 1640 (Life Technologies Inc.) supplemented with 10% FBS. CFPAC-1 and HCT116 cell lines were cultured in Iscove’s Modified Dulbecco’s Medium (Life Technologies Inc.) and McCoy’s 5A Modified Medium (Life Technologies Inc.) supplemented with 10% FBS, respectively. Panc04_03 and Panc02_03 (from Troy Ketela, Donnelly Centre and Banting & Best Department of Medical Research, Toronto, ON) were cultured in RPMI 1640 supplemented with 10 Units/ml human insulin (85%) and 15% FBS. NMuMG cells were transfected using Effectene (QIAGEN) according to the manufacturer’s instructions. Stable HEK 293T, PANC-1, HPAF-II, BxPC3, A549, H727 (human) and NMuMG (murine) ARHGEF2 knockdown cell lines were established by co-transfecting the packaging cell line HEK 293T with human or murine ARHGEF2 lentiviral hairpin plasmids and packaging plasmids pPAX2 and VSV-g using the CalPhos Mammalian Transfection Kit (Clontech). Lentiviral supernatants were collected, filtered and incubated with the target cells in the presence of 8g/ml Polybrene (Sigma). After 48h cells were subjected to puromycin (Sigma) selection (6g/ml for PANC-1, 3g/ml for HPAF-II, 2g/ml for HEK 293T and BxPC3 and 4g/ml for NMuMG cells) until all untransduced cells died. For proliferation assays, Panc04_03, Panc02_03 and PL-45 cells were selected with 2.5g/ml, 2g/ml and 4g/ml puromycin, respectively. All cultures were maintained in a 5% CO2 environment at 37oC.

Expression constructs: Full-length (Arhgef2 or Arhgef21-985) and mutated (Arhgef2T247F) versions of murine and human ARHGEF2 cDNA (accession no. AF177032 and NM_004723.3, respectively) were subcloned into the pFlag-CMV2 vector (Sigma) or pEGFP-C1 (Invitrogen). Full-length murine p115RhoGEF cDNA (accession no. NM_001130150.1) was subcloned into pFlag-CMV2 vector. Murine and human ARHGEF2 pLKO.1 lentiviral shRNA constructs are as

68 described in Chapter 2 (Section 2.3, Table 1). pCDNA3-Pyo-KSR-1 wild-type and mutant expression vectors were kind gifts from Deborah Morrison and were generated as described in Muller et al., 2001.

Cell treatments: For MEK and PI3K inhibition experiments OV-90, CFPAC-1, SK-OV-3, HCT116 or PANC-1 cell lines were cultured in full medium and incubated with PD98059, UO126 or LY294002 (Sigma) diluted in DMSO (Sigma) for 48h. For ROCK inhibition experiments, NMuMG cells were cultured in full medium and treated with 10M Y27632 (Sigma) for 48h. For TGF induction experiments, human TGF1 (Cell Signaling, CN 8915) was diluted in 20M Citrate at a pH of 3.0 to a stock concentration of 100g/ml. TGF1 was added to complete cell culture medium at a final concentration of 10nm/l over indicated time periods and was replaced every 24h.

Western blotting: Cells were scraped into ice-cold lysis buffer (30mM Tris pH7.5, 150mM NaCl,

1% Triton X-100, 0.2% sodium deoxycholate, 10mM NaF, 1mM Na3VO4 and 1mM PMSF) with Complete Protease Inhibitor cocktail (Roche) and incubated on ice for 20min, followed by centrifugation at 16,060xg at 4oC for 10min. Cleared lysates were resuspended in 2X sample buffer, boiled and protein resolved by SDS-PAGE before transfer to PVDF membranes and immunoblotting. For experiments analysing epithelial cell marker expression, cells were lysed directly in 2X sample buffer containing 62.5mM Tris-HCl pH 6.8, 2.5% SDS, 10% glycerol, 5% -mercaptoethanol and 0.02% bromophenol blue, boiled and resolved by SDS-PAGE as described above.

Biotechnology, V9, SC 6260), anti-Zeb1 (Santa Cruz Biotechnology, H-102, SC 25388), anti- alpha tubulin (Molecular Probes), anti-actin (Sigma), anti-GAPDH (Invitrogen), anti-Flag (M2, F3165, Sigma) and anti-GFP (Invitrogen, G10362). HRP-conjugated anti-mouse or anti-rabbit secondary antibodies were from GE Healthcare.

Quantitative real-time reverse transcription PCR: Total cellular RNA was extracted from PANC-1shGFP or PANC-1shGEF1 cell lines using the RNeasy Plus Mini Kit (QIAGEN). 1g of RNA was converted into double-stranded cDNA at 42oC using ImProm-IITM Reverse Transcription System (Promega, Madison, USA) following the manufacturer’s instructions. Quantitative PCR was performed with 50ng of template cDNA mixture from each cell line with SYBR Green human ARHGEF2, CDH1 and GAPDH primers (Primer Bank) (Table 2).

Table 3.1: Gene Target Primer Sequences

GENE Forward primer Reverse primer ARHGEF2 5'-CAGGCATGACCATGTGCTATG- 5’-TTTACAGCGGTTGTGGATAGTC- 3' 3’ CDH1 5’- CGAGAGCTACACGTTCACGG- 5’- GGGTGTCGAGGGAAAAATAGG- 3’ 3’ GAPDH 5’-ACCACAGTCCATGCCATCAC-3’ 5’-TCCACCACCCTGTTGCTGT-3’

Quantitative PCR was performed using the CFX96 Real-Time PCR Detection System from Bio- Rad, and results were analyzed using the software Bio-Rad CFX. Gene expression levels in the samples were calculated relative to control using the comparative CT method: CT = CTsample – -CT CTcontrol, fold change = 2 . GAPDH expression was used to normalize target gene expression levels.

Proliferation assays: Panc02_03, Panc04_03 and PL-45 cells were plated in the appropriate growth medium containing 8ug/ml polybrene and infected with lentiviral expression contructs encoding two distinct shRNAs targeting human ARHGEF2 (shGEF1 and shGEF2), shGFP (negative control) or shPSMD1 (positive control). After 24h the cell culture medium was replaced with fresh medium containing puromycin. 48h later infected cells were harvested by

70 trypsinization, counted and re-plated at 3000 cells/well on a 96-well plate in quadruplicate for phenotypic analysis. 72h later cells were washed with 1X PBS, fixed in 4% PFA, permeabilized with 0.1% Triton X-100 and stained with Hoechst. Nuclei were imaged using a GE IN Cell Analyzer 2000 and nuclei were counted using IN Cell Developer Toolbox 1.9 software. Results were plotted as percent proliferation relative to the shGFP control-expressing cells.

BrdU incorporation: PANC-1-shGFP, PANC-1-shGEF1 and PANC-1-shGEF2 stable cell lines were plated at 1x103 cells per well in a 96-well microplate in quadruplicate. BrdU reagent (Roche) was added to cells after 24h and incorporation was measured after 24h by colorimetric detection as per the manufacturer’s protocol (Roche, 11647229001). Values reflect percentage BrdU incorporation relative to shGFP-expressing cells and represent the mean of three independent experiments.

Immunofluorescence imaging: Cells grown on glass coverslips were treated as indicated in the corresponding figure legends and fixed with 4% PFA for ten minutes, washed three times with 1X PBS and permeabilized with 0.1% Triton X-100 for 5min. The coverslips were blocked with 0.5% w/v BSA in 1X PBS for 1h at room temperature and incubated with primary antibody (anti-rabbit anti-E-cadherin 1:100, anti-mouse anti-vimentin 1:50) in 0.5% BSA/1X PBS at 37oC for 30 min or at 4oC overnight. Coverslips were washed three times with 1X PBS and incubated with secondary antibody (1:500) at 37oC for 1h. Slides were washed once with 1X PBS, incubated with DAPI stain (Invitrogen) at a final concentration of 1:30000 in PBS for 5min at 20oC and washed a final time with 1X PBS. Slides were mounted using GelTol mounting medium (Shandon Immunon, Thermo Electron Corporation). Confocal imaging was performed with an Olympus IX81 inverted microscope using a 60X zoom x3(1.4 NA; PlanApo, Nikon) objective, and FluoView software (Olympus, Tokyo, Japan). Resolution was 512x512 with 12 bits/pixel. The following excitation wavelengths were used for green (473 nm), Texas Red (559 nm) and blue (358nm). All images in each set of experiments were acquired with the same microscope sensitivity settings. All images compared within each figure panel were acquired on the same day, with identical staining conditions, gain and contrast settings, and the same magnification.

Animal studies: All animal studies were carried out using protocols that have been approved by the UHN Animal Care Committee. Xenograft studies in severe combined immunodeficient

(SCID) mice with PANC-1, HPAF-II and BxPC3 cell lines were performed with 2x105 cells resuspended in serum-free medium and injected subcutaneously into the abdominal tissue of the mice. The mice were kept for up to 3 months and tumor measurements were taken bi-weekly. When tumors reached a diameter of 1.5cm or became ulcerated, the mice were sacrificed by carbon dioxide asphyxiation. The tumors were removed, weighed, measured and fixed in 10% buffered formalin for histologic processing or flash-frozen in liquid nitrogen for protein and/or RNA analysis. 5 injections were performed per condition and each cell line was performed in duplicate. Tumor measurements were taken with a calliper and tumor volume was calculated by the ellipsoid formula V=/6 x (l x w2), where l and w denote the longest and shortest diameter, respectively.

Immunohistochemistry: For human pancreatic tissue analysis, tissue microarrays (TMA) of normal pancreatic ducts, pancreatic intraepithelial neoplasia (PanIN) lesions and adenocarcinoma were constructed from paraffin blocks of Whipple resection specimens, as described previously (Al-Aynati et al., 2004), and following a study protocol approved by the UHN Research Ethics Board. TMAs from normal colon, primary and metastatic colorectal tumors or from normal lung and primary lung adenocarcinomas (ADC) harboring wild-type K-RAS or K-RASD12 mutations were similarly derived. Immunohistochemistry was performed using the Biotin-Streptavidin- HRP detection system and a human Arhgef2 antibody (14B11 mouse monoclonal antibody) at a 1:500 dilution. To evaluate the expression levels of Arhgef2, staining intensity in the ductal, epithelial cells or lesions were judged by two pathologists and scored as 3 (strong staining), 2 (moderate staining) or 1 (weak staining). As staining was observed to be diffuse in the tumors analyzed, percentage of tumor cell stained was not recorded. Tumor sections derived from PANC-1shGFP and shGEF1 xenografts were derived as described under Animal Studies and were probed for caspase 3 cleavage using anti-cleaved caspase 3 (Asp 175) antibody (CST, 9661) using the Biotin-Streptavidin-HRP detection system.

RNA preparation and microarray: Total RNA was isolated from cultured cells using the RNeasy Mini kit (QIAGEN). The quality of RNA was verified using agarose gel and the Agilent bioanalyzer (Agilent technologies, Palo Alto, CA). 200ng of RNA were labeled using Illumina TotalPrep-96 RNA Amplification kit (Ambion, lot 1107026) as per the amplification protocol. 750ng of cRNA (PANC-1shGFP and PANC-1shArhgef2 cell lines) generated from amplification

72 and labeling were hybridized into 1 Human HT-12 v4.0 BeadChip and 1.5ug cRNA (NIH 3T3- H-RASV12shGFP and NIH 3T3-H-RASV12shArhgef2 cell lines) was used for mouse WG-6 v2. The BeadChip was incubated at 58oC at rotation speed 5 for 18h for the hybridization. The BeadChips were washed and stained as per the Illumina protocol and scanned on the iScan (Illumina). The data files were quantified in GenomeStudio Version 2011.1 (Illumina). All samples passed Illumina’s sample dependent and independent QC metrics. Sample preparation and hybridization were done at the UHN Microarray Centre at the MaRS Centre (Toronto, Ontario, Canada). Comparative analysis was performed between PANC-1shGFP and PANC- 1shArhgef2 and NIH 3T3-H-RASV12shGFP and NIH 3T3-H-RASV12shArhgef2 cell lines. Functional annotation of gene sets was performed using the DAVID Bioinformatics resource website (http://david.abcc.ncifcrf.gov/) Functional Annotation tool. In PANC-1shGFP vs PANC-

1shArhgef2 cell lines, the 416 most downnregulated genes (<-1.52fold change or <-0.6 DF(log2)) and 399 most upregulated genes (>1.52 fold change or >0.6 DF(log2)) relative to PANC-1shGFP cells were selected for analysis by Biological Process, Molecular Function and KEGG Pathway. For NIH 3T3-H-RASV12shGFP and NIH 3T3-H-RASV12shArhgef2 cell lines, Significance Analysis of Microarrays (SAM) was performed on differentially regulated genes in triplicate RNA samples from each cell line. Thirty-three upregulated and 170 downregulated genes were found to be statistically significant across all samples (p<0.05).

EMT induction: NMuMG or PANC-1 cells were infected with LVGFP and/or LVArhgef2 and selected in puromycin-containing medium for 3 days prior to TGF treatment. For rescue experiments, cells were transfected with expression plasmids 16h prior to TGF treatment. NMuMG cells were trypsinized and plated in 6-well plates to achieve 60-80% confluence the day of treatment. Cells were treated with DMEM supplemented with 10% FBS and containing 10ng/ml TGF1; medium was refreshed every 24h. For immunofluorescence studies, NMuMG cells were plated directly on coverslips in a 6-well format and treatments carried out as described under Immunofluorescence studies.

Statistical analyses: Values are expressed as means +/- SD or +/- SE as indicated. Paired Student’s t-tests (Kirkman, 2006) were performed to determine statistical significance between samples. Experiments were performed at least three times and means with p < 0.05 were considered statistically significant.

3.4 Results

3.4.1 ARHGEF2 is essential across several human epithelial cancer cell lines and its protein expression is regulated by the RAS/MAPK pathway

Considering the role of Arhgef2 in RAS-mediated cell survival in fibroblast cells, we sought to determine if Arhgef2 was important for cell viability in human epithelial adenocarcinoma cell lines exhibiting more varied mutatomes. To that end, we analyzed publicly-available gene essentiality data derived from a genome-wide pooled shRNA screen designed to distinguish genes that are essential for cancer cell survival in human breast, colon, lung, ovarian and pancreatic cell lines (Marcotte et al., 2012). ARHGEF2 was found to be highly essential (p < 0.05) across 13 of 72 cell lines, including 4 ovarian, 5 pancreatic, 2 breast, 1 colon and 1 lung cell line (Figure 3.2A). For further validation, we selected four cell lines that displayed the highest statistical significance for ARHGEF2 essentiality. We stably infected these cells with an ARHGEF2 hairpin distinct from those contained in The RNAi Consortium (TRC) screening pool and found that cells depleted of Arhgef2 protein exhibited increased cell death relative to hairpin control-expressing cells (Figures 3.2B and 3.2C). These data suggest that Arhgef2 is essential for cell survival in human cell lines derived from different tumor types.

Of the 13 proposed ARHGEF2-essential cell lines, 8 have gain-of-function mutations in components of the Ras/MAPK pathway, including H-RAS (Hs578T), K-RAS (CFPAC-1, HCT116, Panc02_03, PaTu_8988T, IMIM-PC-1, RWP-1) and B-Raf (OV-90), suggesting that ARHGEF2 essentiality has a preference for cells with elevated MAPK activity (Hollestelle et al., 2007, Moore et al., 2001, Shirasawa et al., 1993, Jaffee et al., 1998, Shen et al., 2008, Estep et al., 2007). To assess whether Arhgef2 expression was dependent on MAPK activation, we treated OV-90, CFPAC-1, SK-OV-3 and HCT116 cells with two MEK1/2 inhibitors, PD98059 and UO126, and found that Arhgef2 protein expression decreased with MEK1/2 inhibition (Figure 3.3). These data demonstrate that Arhgef2 protein expression is regulated by the MAPK pathway in human epithelial cell models bearing endogenous mutations in the Ras/MAPK pathway and is a critical mediator of cell survival in these cells, in agreement with our studies in murine fibroblast cells.

Figure 3.2 ARHGEF2 is essential across several human epithelial cancer cell lines. (A) 72 breast, ovarian and pancreatic cell lines were infected with 78,432 shRNAs targeting 16,056 genes for an average of 5 shRNAs per gene (for method see Moffat et al., 2012). Gene dropout signatures were determined by calculating the shRNA Rank Profile (shARP) of each gene. Gene essential shARP scores for ARHGEF2 were significant (p<0.05) in 13 cell lines, including 2 (13.3%) breast, 1 (6.6%) colon), 1 (6.6%) lung, 4 (26.6%) ovarian and 5 (33.3%) pancreatic. P- values for each cell line are depicted schematically in order of significance and cancer cell types are grouped by color. (B) Representative cell densities of 4 ARHGEF2-essential cell lines 6 days following infection with a hairpin control (shGFP) or a lentiviral shRNA targeting human ARHGEF2 (shGEF). (C) Western blot analysis validating Arhgef2 protein knockdown in HEK 293T cells expressing shGFP or shGEF with tubulin serving as a protein loading control.

3.4.2 Arhgef2 is required for PDAC tumor growth in vivo

We sought to determine whether Arhgef2 was required for tumor growth in human epithelial cell models harboring endogenous RAS mutations. Over 95% of human pancreatic cancers harbor

Figure 3.3 ARHGEF2 protein expression is regulated by the RAS/MAPK pathway in human epithelial cell lines. Immunoblot analysis of Arhgef2 expression after 48h of DMSO or MEK inhibitor treatment with PD98059 (30M) or UO126 (10M) in 4 ARHGEF2-essential cell lines. Activating phosphorylations of ERK1/2 kinases indicate MEK1/2 inhibition and total ERK1/2 protein levels are shown as gel loading controls. Data are representative of two independent experiments. endogenous activating mutations in K-RAS, making them ideal model systems to study the mechanisms contributing to aberrant RAS signaling (Smit et al., 1988, Almoguera et al., 1988, Grunewald et al., 1989). Thus, we examined the requirement of Arhgef2 for pancreatic tumor growth of three PDAC cell lines, PANC-1 (K-RASD12), HPAF-II (K-RASD12) and BxPC3 (wild- type K-RAS) in immunodeficient mice (Moore et al., 2001, Aoki et al., 1997). Arhgef2 was knocked down in each of these cell lines using two distinct hairpins and Arhgef2 protein depletion was confirmed by immunoblotting (Figures 3.4A, 3.4C and 3.4E, insets). PANC-1 and HPAF-II cells exhibited a 90% (PANC-1) and 70% (HPAF-II) decrease in mean tumor volume and weight relative to hairpin control cells (Figures 3.4A-D) with increased tumor-associated caspase 3 cleavage in PANC-1 cells as assessed by immunostaining (Figure 3.4G). By distinction, tumor growth of the K-RAS wild-type BxPC3 cells showed no dependence on Arhgef2 expression (Figures 3.4E and 3.4F). These data show that cell lines bearing activating mutations in RAS require Arhgef2 expression for tumor growth.

We sought to determine the role of Arhgef2 for KSR-1-MAPK activation in pancreatic adenocarcinoma cells by monitoring the phosphorylation state of KSR-1 and ERK1/2 in Arhgef2 knockdown PANC-1 cells. In cells stably depleted of Arhgef2, we observed increased phosphorylation of KSR-1 on S392 and a corresponding decrease in phosphorylation of ERK1/2 compared to hairpin control-expressing cells (Figure 3.5A). Expression of an shRNA-resistant active form of Arhgef2 (rArhgef287-151) or KSR-1S392A, but not wild-type KSR-1, restored basal levels of phosphorylated KSR-1 and ERK1/2 in Arhgef2-depleted PANC-1 cells. These data

76 indicate that Arhgef2 is both necessary and sufficient for KSR-1 S392 dephosphorylation and subsequent ERK1/2 activation in pancreatic cells harboring mutations in endogenous RAS.

Figure 3.4 Arhgef2 is required for pancreatic tumor growth in vivo. (A-F) PANC-1, HPAF-II and BxPC3 cells were infected with a hairpin control (shGFP) or two distinct hairpins targeting human ARHGEF2 (shGEF1 and shGEF2). Arhgef2 protein expression was assayed by Western blot and tubulin served as a protein loading control (A, C, E, inset). 2x105 shGFP, shGEF1 and shGEF2 cells were injected subcutaneously into SCID mice and allowed to form xenografts over the indicated time periods (growth curves depicted in A, C and E). Tumors were harvested once control tumors reached a diameter of 1.5cm. Final tumor volumes were determined for each cell line and are depicted graphically in B, D and F. Representative images of dissected tumors are shown (below bar graphs) from one of two experiments performed per cell line. Error bars represent SD of one representative experiment from n=5 tumors (**p<0.01, *p<0.05). (G) Representative immunohistochemical images of xenografts derived from shGFP or shGEF2-expressing PANC-1 cells stained for cleaved caspase 3. Cleaved caspase 3 expression is depicted in brown. 40 images derived from 5 PANC-1-shGFP tumors and 40 images 5 PANC-1-shGEF2 tumors were analyzed.

Furthermore, knockdown of Arhgef2 in PANC-1, Panc02_03, Panc05_04 and PL-45 PDAC cell lines resulted in a 30-70% decrease in cell growth relative to hairpin control-expressing cells (Figures 3.5B and 3.5C). These data show that Arhgef2 affects cell growth in several PDAC cell line models harboring mutant RAS and demonstrate that the requirement of Arhgef2 for RAS/MAPK signaling is conserved across these cell types.

Figure 3.5 Arhgef2 is required for KSR-1 S392 dephosphorylation, ERK1/2 phosphorylation and proliferation in PDAC cells. (A) PANC-1 cells were infected with a hairpin control (shGFP, lane 1) or hairpin targeting human ARHGEF2 (shGEF, lanes 2-5). Arhgef2-depleted cells were subsequently transfected with Flag- Arhgef287-151 (lane 3), Pyo-KSR-1S392A (lane 4) or wild-type Pyo-KSR-1 (lane 5). Total cell lysates were probed for ERK1/2 phosphorylation and KSR-1 S392 phosphorylation (KSR-1 pS392) by immunoblotting (rows 3 and 4). Arhgef2 expression was probed using endogenous Arhgef2 antibody and Pyo-KSR-1 was detected using anti-KSR-1 antibody. Levels of RAS and ERK1/2 were probed to control for total protein levels. (B) PANC-1 cells stably expressing shGFP, shGEF1 or shGEF2 were plated at 1x103 cells/well in quadruplicate in a 96-well plate and BrdU incorporation was measured after 24h by colorimetric detection. Data are depicted as percent BrdU incorporation compared to shGFP-expressing cells and are the mean of three independent experiments +/- SE. (C) Panc04_03, Panc02_03 and PL-45 cells were infected with shGFP (negative control), shPSMD1 (positive control) or two distinct hairpins targeting human ARHGEF2 (shGEF1 and shGEF2). 3x103 hairpin-expressing cells were plated in quadruplicate in 96-well plates and cell number was determined after 72h by nuclei staining. Data are represented as percent proliferation relative to shGFP-expressing cells +/- SD.

3.4.3 Arhgef2 expression correlates with advanced tumor grade in human lung, colorectal and pancreatic cancer

K-RAS mutations most frequently occur in PDAC, colorectal and NSCLC tumors (Almoguera et al., 1988, Moskaluk et al., 1997, Andreyev et al., 2001, Bardelli et al., 2005). We sought to evaluate whether Arhgef2 expression was increased in these tumor types by performing immunohistochemistry on lung, colorectal and pancreatic tissue microarrays (TMAs) (Figure 3.6A-C). Normal lung TMAs expressed weak immunostaining for Arhgef2 that was significantly increased in primary adenocarcinoma (ADC) and further enhanced in ADCs harboring mutations in K-RAS (Figure 3.6A). Of 63 normal lung sections analyzed, 49.21% (31/63), 34.92% (22/63) and 15.87% (10/63) stained weakly, moderately, and strongly for Arhgef2, respectively (Figure 3.6A, side graph). By contrast, 11.11% (7/63), 36.51% (23/63) and 53.97% (34/63) primary ADCs stained weakly, moderately, and strongly for Arhgef2. A similar difference in expression trends was observed in large cell undifferentiated carcinomas of the lung (LCUL) (71.43% (5/7) vs 14.3% (1/7) weakly staining, 28.57% (2/7) vs 28.57% (2/7) moderately staining and 0% (0/7) vs 57.14% (4/7) strongly staining for normal and LCUL samples, respectively), while the differences were less pronounced but still correlative in squamous cell lung carcinomas (SQ) (60% (18/30) vs 36.67% (11/30) weakly staining, 26.67% (8/30) vs 40% (12/30) moderately staining and 13.33% (4/30) vs 23.33% (7/30) strongly staining for normal and SQ samples, respectively). In colorectal TMAs, 62.1% (18/29) of normal tissues were absent for Arhgef2 staining, 31.0% (9/29) showed weak staining and 6.9% (2/29) showed strong staining. By distinction, 19.4% (13/67) primary and 22.9% (8/35) metastatic colon ADCs stained moderately for Arhgef2 while 80.6% (54/67) primary and 77.1% (27/35) metastatic lesions exhibited strong Arhgef2 staining (Figure 3.6B, side graph). Moreover, analysis of 14 normal pancreatic ducts, 32 PanIN1 (A and B) lesions, 9 PanIN2 and I3 lesions and 14 advanced PDAC samples for Arhgef2 expression revealed that in all normal pancreatic ducts, PanIN1 and PanIN2 lesions exhibited weak Arhgef2 immunostaining (Figure 3.6C). 77.8% (7/9) pre-invasive PanIN2 and 3 lesions showed moderate to strong Arhgef2 staining, with the majority (71.4%, 5 of 7) showing lesser intensity than those observed in the majority of advanced PDAC cases (Figure 3.6C, side graph). All 14 PDAC cases stained for Arhgef2, with 71.4% (10/14) exhibiting strong staining. These data demonstrate that Arhgef2 expression is elevated in primary RAS-mutated tumors and is

79 increased with advanced tumor grade, suggesting that Arhgef2 may play a role in more advanced stages of tumorigenesis.

3.4.4 Arhgef2 expression alters gene signatures associated with epithelial differentiation state

Given that Arhgef2 expression is increased with advanced tumor grade in RAS-mutated cancers, we addressed if Arhgef2 may contribute to the acquisition of migratory phenotypes associated with late stages of tumorigenesis and EMT. A key feature of cells that have undergone EMT is the acquisition of new transcriptional programs to maintain their mesenchymal phenotypes (Radisky, 2005). Mesenchymal gene expression signatures have been used to distinguish tumor subtypes and often correlate with even poorer overall survival (Kim et al., 2010, Hoshida et al., 2009).

In order to gain broader insight into ARHGEF2-regulated gene signatures in epithelial tumor cells, we compared gene expression profiles of PANC-1 cells stably expressing a hairpin control to those expressing an ARHGEF2 shRNA. Gene ontology analysis of the 399 most upregulated genes in ARHGEF2-depleted PANC-1 cells revealed that biological adhesion/cell adhesion (p<0.00005) and cell motion/migration (p=0.0012) were among the biological processes most significantly perturbed, according to the DAVID algorithm (Dennis et al., 2003) (See Appendix, Table IA). Categorizing genes by cellular component showed that there was a significant alteration in genes localized to cell junctions (Appendix, Table IB, p=0.0011). KEGG pathway analysis further confirmed these trends, identifying an enrichment in genes regulating focal adhesions (p=0.0003) and ECM-receptor interactions (p=0.0077) (Appendix, Table ID). Among

Figure 3.6 Arhgef2 expression correlates with advanced tumor grade in human lung, colon and pancreatic tissue microarrays. (A) Representative images of lung TMAs showing Arhgef2 protein expression in normal lung tissue (first panel), a lung adenocarcinoma (ADC) expressing wild-type K-RAS (second panel) and a lung ADC with a K-RASD12 mutation. Arhgef2 expression was detected with a monoclonal antibody against human Arhgef2 and is depicted in brown. Staining was scored as weak (1), moderate (2) or strong (3) by two pathologists. The distribution of Arhgef2 immunoscores across tumor groups is shown in the adjacent bar graph. (B) TMAs showing representative Arhgef2 protein expression in the normal colon (first panel), a primary colorectal tumor (second panel) and a metastatic colorectal lesion (third panel). 29 normal tissues, 67 primary and 35 metastatic colorectal tumors were stained for Arhgef2 and quantified as in (A). (C) Arhgef2 protein expression in PDAC. Representative images of TMAs of pancreatic ducts (normal), pancreatic intraepithelial neoplasia (PanIN-1B and PanIN-3) lesions and adenocarcinoma (ADC) probed for Arhgef2 expression. Staining intensity in the ductal cells or lesions was

scored as described in (A). Distribution of Arhgef2 immunoscores for 14 normal, 18 PanIN-1A, 14 PanIN-1B, 3 PanIN-2, 6 PanIN-3 and 14 ADC TMA samples is shown graphically (right). the genes found to be induced were genes encoding E-cadherin, claudins 1,10 and 12, integrins alpha 2, 5 and beta 1, laminin gamma 2, collagens type I, alpha 1, type V, alpha 2, and type XI, alpha 1 and PAK1 and 6 proteins (Appendix, Tables IIB-F). Functional annotation of the 416 most downregulated genes in ARHGEF2-depleted PANC-1 cells, in turn, showed that mesenchymal differentiation/development was one of the most significant biological processes perturbed (Appendix, Table IIIA, p<0.00005). Significant gene enrichment in focal adhesion (p=0.0288) and adherens junction (p=0.0470) cellular components were also identified (Appendix, Table IIIB). KEGG pathway analysis of the gene set showed that genes involved in the regulation of focal adhesions was also significantly altered (Appendix, Table III, p=0.0209). The genes associated with each process are listed in Table IV. These data demonstrate that stable depletion of ARHGEF2 results in the perturbation of epithelial adhesion, motility and differentiation gene signatures, suggesting that Arhgef2 may functionally contribute to these biological processes in vivo.

In order to probe the effect of ARHGEF2 on established mesenchymal gene signatures, we performed microarray on RASV12-transformed fibroblast cells stably depleted of ARHGEF2 used in our previous studies (Appendix, Table VA). Significance Analysis of Microarrays (SAM) revealed 202 genes whose expression was significantly perturbed following ARHGEF2 knockdown (31 upregulated, 171 downregulated). Gene ontology analysis of the downregulated gene set according to biological process showed that a subset of genes regulating epithelial cell differentiation were suppressed, including fibroblast growth factor 10 (FGF10), fibroblast growth factor receptor 2 (FGFR2), keratin 14 (KRT14) and transforming growth factor beta 1 (TGF1) (p=0.0101) (Appendix, Tables VA and VIB). Pathways controlling motility and migration were also downregulated, including response to wounding, chemotaxis and the regulation of morphogenesis (p=0.0026, p=0.0066 and p=0.0126, respectively) (Appendix, Table VA). Grouping genes by cellular component showed that genes involved in the regulation of adherens junctions, anchoring junctions, cell junctions and focal adhesions were most significantly suppressed following depletion of ARHGEF2 (p=0.0007, p=0.0014, p=0.0042 and p=0.0066, respectively) (Appendix, Table VIB). Likewise, KEGG Pathway analysis revealed a significant concentration of genes regulated by ARHGEF2 involved in tight junction formation (p=0.0056)

(Appendix, Table VID). Together, these data demonstrate that ARHGEF2 depletion results in the upregulation of epitheloid genes in PDAC cells and the downregulation of mesenchymal genes in RASV12-transformed fibroblast cells and suggests that Arhgef2 may functionally contribute to both the induction and maintenance of a mesenchymal cell phenotype.

3.4.5 Arhgef2 suppresses the epithelial cell phenotype in RAS-independent human adenocarcinoma cell lines

We found a number of genes positively associated with epithelial differentiation state upregulated in Arhgef2-depleted PANC-1 cells, including the epithelial cell adhesion molecule EPCAM (2.41-fold change), the tight junction-associated membrane proteins CLDN10 and CLDN12 (2.33 and 1.52-fold changes, respectively), alpha and beta subunits of the adhesion- mediating integrin transmembrane receptors ITG2, ITG5 and ITG1 (2.30, 1.78 and 1.70-fold changes, respectively), the intercellular adhesion-promoting transmembrane receptors TSPAN3, 13 and 18 (2.21, 2.01 and 2.15 fold-change, respectively), the laminin gamma 2 chain LAMC2 (1.91 fold-change), a component of anchoring filaments that connect epithelial cells to the basement membrane, the intermediate filament protein KRT19 (1.73 fold-change) and the adhesion-promoting receptor tyrosine kinase EPHB2 (1.68 fold-change) (Kubota et al., 1999, Witkowski et al., 1993, Yanez-Mo et al., 2001, Pakkala et al., 2002, Pfaff et al., 2008) (Figure 3.7A). The positive role for many of these molecules in the regulation of epithelial cell integrity is supported by the observation that their downregulation has been implicated in increased migration, invasion and metastasis in human tumors, as has been found with the claudin, laminin, integrin and ephrin families of proteins (Ip et al., 2007, Karamitropoulou et al., 2011, Witkowski et al., 1993, Dong Li Guo et al., 2005).

The loss of the epithelial adhesion molecule E-cadherin is an essential step regulating the dissemination of cell-cell junctions and promoting EMT (Lehembre et al., 2008). We found that E-cadherin mRNA was elevated by 2.2-fold in ARHGEF2-depleted PANC-1 cells by microarray, suggesting that Arhgef2 may positively regulate EMT by suppressing E-cadherin transcription (Figure 3.7A). To confirm that E-cadherin was upregulated in epithelial cells lacking ARHGEF2 expression, we performed quantitative PCR on PANC-1 cells expressing a

83 hairpin control or an ARHGEF2 shRNA (Figure 3.7B). We found that E-cadherin transcripts were elevated by over 18-fold in ARHGEF2-depleted cells relative to GAPDH expression (p=0.0099). Immunoblot analysis of PANC-1 cells harboring stable Arhgef2 knockdown confirmed that the elevation of E-cadherin expression was maintained at the protein level (Figure 3.7C). Furthermore, we found a slight decrease in expression of the mesenchymal marker vimentin (Figure 3.7C). Together, these results demonstrate that Arhgef2 regulates the expression of genes associated with epithelial differentiation state.

Figure 3.7 Arhgef2 suppresses the epithelial cell phenotype in human adenocarcinoma cell lines. (A) List of genes associated with epithelial differentiation state shown to be upregulated by at least 1.52-fold (DF log2 of > 0.6) in ARHGEF2-depleted PANC-1 cells relative to shGFP-expressing cells by microarray analysis. Fold change is indicated for each gene. (B) Validation of E-cadherin gene (CDH1) upregulation by real-time quantitative PCR. Transcript levels of ARHGEF2 and CDH1 were normalized to GAPDH expression and are represented as fold decrease and increase of PANC-1shGEF over PANC-1shGFP-expressing cells, respectively. Data are the mean of four independent experiments for ARHGEF2 (p1=0.011) and two independent experiments for CDH1 (p2=0.0099) +/- SE. (C) Validation of E-cadherin protein upregulation in Arhgef2-depleted PANC-1 cells compared to shGFP- expressing cells by Western blot. Arhgef2 expression shows level of knockdown and vimentin expression is shown as a mesenchymal marker, with total ERK1/2 serving as a protein loading control. E-cadherin is functional when it is localized to cell-cell junctions of epithelial cells, where it regulates cell adhesion and polarization. We therefore assessed the localization of E-cadherin in Arhgef2-depleted cells by immunofluorescence staining (Figure 3.7D). PANC-1 cells expressing a non-targeting hairpin exhibited low levels of E-cadherin expression that was localized diffusely throughout the cytoplasm (Figure 3.7D, row 1 column 1). In contrast, the cells expressed high levels of vimentin, in agreement with previous reports that PANC-1 cells represent a poorly

84 differentiated PDAC, exhibiting mesenchymal-like properties (Singh et al., 2009) (Figure 3.7D, row 1 column 3, arrowhead). In PANC-1 cells lacking Arhgef2 expression, however, E-cadherin staining was prominently localized to the cell periphery and cells adopted a rounder, cobblestone-like morphology characteristic of epithelial cells (Figure 3.7D, row 2 column 1, arrowhead). Vimentin expression was reduced and localized diffusely throughout the cytoplasm, in contrast to the pools of vimentin seen at specific subcellular sites in hairpin control-expressing PANC-1 cells (Figure 3.7D, row 1 column 3 vs row 2 column 3, arrowhead). Higher magnification images of individual hairpin control and Arhgef2 shRNA-expressing cells are shown in second and third rows, respectively. These data show that Arhgef2 expression contributes to the mesenchymal properties of PANC-1 cells by modulating both the expression and proper localization of E-cadherin and vimentin.

We sought to determine if Arhgef2 suppressed E-cadherin expression in other human adenocarcinoma cell lines exhibiting mesenchymal-like properties. To that end, we compared Arhgef2-dependent changes in E-cadherin and vimentin expression in four cell lines categorized as epithelioid or mesenchymal-like (Singh et al., 2009). These included the mesenchymal-like A549 (lung ADC) and PANC-1 (PDAC) cell lines and the epithelial-like H727 (lung ADC) and HPAF-II (PDAC) cell lines. Biochemical analysis of lysates derived from each cell line confirmed that H727 and HPAF-II cells expressed high levels of E-cadherin and low levels of vimentin, while A549 and PANC-1 cells expressed low levels of E-cadherin and high levels of vimentin, consistent with their previously characterized differentiation states (Singh et al., 2009) (Figure 3.7E). Moreover, low levels of E-cadherin were associated with Zeb1 expression, a transcriptional repressor of E-cadherin (Figure 3.7E) (Sanchez-Tillo et al., 2011). Stable depletion of Arhgef2 in A549 cells potently induced the expression of E-cadherin with two distinct Arhgef2 shRNAs, as was observed in PANC-1 cells (Figures 3.7F and 3.7C, respectively). Importantly, in H727 and HPAF-II cells already expressing E-cadherin, there was no further induction of E-cadherin protein levels, although a decrease in vimentin expressed was observed in the HPAF-II cell line (Figures 3.7G and 3.7H, respectively). Together, these studies demonstrate that Arhgef2 promotes the mesenchymal phenotype of human epithelial cancer cells through the downregulation of E-cadherin expression or the induction of vimentin expression depending on the stage of mesenchymal differentiation of adenocarcinoma cell lines.

Figure 3.7 Arhgef2 suppresses the epithelial cell phenotype in human adenocarcinoma cell lines. (D) Immunofluorescence analysis of PANC-1 cells stably expressing shGFP (rows 1 and 3) or Arhgef2 shRNA (shGEF, rows 2 and 4) for endogenous E-cadherin (column 1) or vimentin expression (column 3). Larger magnification images are shown in rows 3 and 4. Arrowheads show areas of increased E-cadherin junctional localization or cytoplasmic vimentin localization. Images are representative of three independent experiments. (E) Western blot analysis of human epithelial tumor cell lines H727, A549 (ADC), HPAF-II and PANC-1 (PDAC) for E-cadherin, vimentin and Zeb1 expression. Arhgef2 expression levels are shown cross cell lines and tubulin serves as a protein loading control. (F-H) A549, H727 and HPAF-11 cells were stably infected with hairpin control (shGFP) or two distinct shRNAs targeting human ARHGEF2 (shGEF1 and shGEF2) and lysates were probed for E-cadherin and/or vimentin expression. Level of Arhgef2 knockdown is indicated by protein expression of endogenous Arhgef2; GAPDH or tubulin serve as protein loading controls.

3.4.6 Arhgef2 is required for TGF-induced epithelial-to-mesenchymal- transition in a mammary epithelial cell model

To directly address the role of Arhgef2 in the regulation of EMT, we employed a murine mammary epithelial cell model, NMuMG, that undergoes mesenchymal transformation in response to TGF. Treatment of NMuMG cells with TGF for 48 hours resulted in the progressive downregulation of E-cadherin and upregulation of vimentin expression (Figure 3.8A) and paralleled the acquisition of an elongated, mesenchymal-like phenotype from an epithelioid morphology (Figure 3.8B). We also observed an increase of Arhgef2 expression after 24 hours of TGF treatment; however, its expression was reduced to basal levels after 48 hours (Figure 3.8A). Immunofluorescence staining of NMuMG cells before and after 48 hours of TGF treatment showed a prominent decrease in the staining intensity and junctional localization of E-cadherin and a simultaneous increase in the intensity of vimentin staining in the cytoplasm, demonstrating that NMuMG cells functionally transform into mesenchymal cells (Figure 3.8C). To assess the requirement of Arhgef2 in TGF-mediated EMT we stably knocked down murine Arhgef2 in native NMuMG cells with two distinct hairpins and treated cells with TGF for 48

Figure 3.8 TGF induces epithelial-to-mesenchymal-transition in a normal mammary gland epithelial cell model. (A) Western blot analysis of NMuMG cells treated with 10ng/ml TGF1 for 24h or 48h. Expression of Arhgef2, E-cadherin and vimentin are shown and GAPDH serves as a protein loading control. (B) Phase-contrast images of NMuMG cells untreated (first panel) or treated with 10ng/ml TGF1 for 24h (panel 2) or 48h. Images are representative of four independent experiments. (C) Immunofluorescence analysis of NMuMG cells treated with TGF1 as in (A) and (B). Cells were fixed and stained for E-cadherin (ECAD, column 1), vimentin (VIM, column 3) or DAPI (columns 2 and 4) after 48h of treatment with vehicle control (first row) or 10ng/ml TGF1 (second row). hours (Figures 3.9A and 3.9B). Although Arhgef2 expression was highly suppressed with both shRNAs, shGEF1 showed greater knockdown efficiency than shGEF2 (Figure 3.9A, fourth row). Since the downstream effector of Arhgef2, ROCK, has shown to be required for TGF-induced EMT in this cell system, we compared the effects of ROCK and Arhgef2 inhibition by co-

treating cells with the ROCK inhibitor Y27632 and TGF (Figure 3.9A, lanes 7 and 8 and Figure 3.9B, column 4) (Bhowmick et al., 2001). Immunoblot analysis for E-cadherin and vimentin

Figure 3.9 Arhgef2 is required for TGF-induced epithelial-to-mesenchymal-transition in NMuMG cells. (A) Western blot analysis of NMuMG cells stably expressing a hairpin control (shGFP) or two distinct shRNAs targeting murine Arhgef2 (shGEF1 and shGEF2) and treated with vehicle control (lanes 1, 3, and 5), 10ng/ml TGF1 (lanes 2, 4 and 6), or 10M of the ROCK inhibitor Y27632 alone or with TGF1 (lanes 7 and 8, respectively) for 48h. Total cell lysates were probed for Arhgef2, E-cadherin and vimentin expression levels and GAPDH serves as a protein loading control. (B) Phase-contrast images of NMuMG cells described in (A). Images are representative of three independent experiments.

showed that Arhgef2 depletion and ROCK inhibition did not prevent the TGF-induced downregulation of E-cadherin expression and did not significantly alter vimentin expression (Figure 3.9A). However, the morphological transition to a mesenchymal phenotype was strongly inhibited in Arhgef2-depleted cells in response to TGF in a manner that correlated with the level of Arhgef2 knockdown (Figure 3.9B, columns 2 and 3). Moreover, ROCK inhibition partially suppressed the development of a fibroblastoid phenotype in response to TGF, although the effect was not as strong compared to Arhgef2 depletion with either hairpin (Figure 3.9B, column 4). Visualization of E-cadherin and vimentin expression by immunofluorescence staining showed that although the intensity of E-cadherin expression decreased in both hairpin control and Arhgef2 hairpin-expressing cells in response to TGF, some peripheral E-cadherin expression was maintained in the absence of Arhgef2 (Figure 3.9C, column 1, rows 2 and 4).

Figure 3.9 Arhgef2 is required for TGF-induced epithelial-to-mesenchymal-transition in NMuMG cells. (C) Immunofluorescence analysis of NMuMG cells stably expressing a hairpin control (shGFP) or Arhgef2 shRNA (shGEF1) treated with vehicle control (rows 1 and 3) or 10ng/ml TGF1 (rows 2 and 4) for 48h. Cells were fixed and stained for E-cadherin (column 1), vimentin (column 3) and DAPI (columns 2 and 4). Furthermore, the cells failed to adopt an elongated, spindle-shaped morphology and instead remained round and flattened. Arhgef2-depleted NMuMG cells exhibited a decrease in vimentin staining intensity in response to TGF compared to hairpin control-expressing cells (Figure 3.9C, column 3, rows 2 and 4). Together, these data show that Arhgef2 is required for the full transition from an epithelial to a mesenchymal cell state in response to TGF.

Figure 3.9 Arhgef2 is required for TGF-induced epithelial-to-mesenchymal-transition in NMuMG cells. (D) Phase-contrast images of NMuMG cells stably expressing shGFP, shGEF1 and shGEF2 and treated with 10ng/ml TGF1 for 48h. Cells were transfected with Arhgef287-151 (column 2) or p115RhoGEF (column 3) 24h prior to TGF1 treatment.

To discern the mechanism by which Arhgef2 is required for TGF-induced EMT, we expressed active Arhgef287-151 and p115RhoGEF in NMuMG cells depleted of Arhgef2 (Figures 3.9D and 3.9E). Expression of either GEF was unable to fully rescue the mesenchymal phenotype induced by TGF; however, both induced similar intermediate fibroblastoid morphologies in an Arhgef2- depleted background (Figure 3.9D, columns 2 and 3 and rows 2 and 3). These data suggest that increased Rho activity can partially compensate for a loss of Arhgef2 expression and that Arhgef2 may promote EMT – at least in part - via its exchange activity on RhoA.

Figure 3.9 Arhgef2 is required for TGF-induced epithelial-to-mesenchymal-transition in NMuMG cells. (E) Western blot analysis showing Arhgef287-151 (upper panel) and p115RhoGEF (lower panel) expression in transfected NMuMG cells described in (D). GAPDH and actin serve as protein loading controls in each respective panel.

3.5 Discussion

Pancreatic ductal adenocarcinoma (PDAC) is one of the most fatal human malignancies, with a 5-year survival rate of less than 5% (Warshaw et al., 1992). Its poor prognosis is largely attributed to the advanced stage of the disease usually presented at the time of diagnosis and the refractory nature of PDAC to current therapies (Warshaw et al., 1992, Jemal et al., 2005). Although mutant K-RAS plays a causal role in pancreatic tumorigenesis, drugs targeting RAS have yet to be clinically effective (Van Cutsem et al., 2004). Thus, the identification of RAS effectors that are essential for tumor cell survival is critical in order to improve treatment strategies against this disease.

In this chapter we found that Arhgef2 is required for cell survival and transformation in RAS- mutated human epithelial pancreatic xenograft models and contributes to RAS signaling in these cells by a parallel mechanism to that observed in fibroblasts. Furthermore, analysis of Arhgef2 expression at progressive stages of human lung, colon and pancreatic tumorigenesis revealed that Arhgef2 levels are dramatically increased with advanced tumor grade. Increased Arhgef2 expression promotes the genetic and morphological transition from an epithelial to a mesenchymal cell phenotype and maintains low E-cadherin levels in mesenchymal-like mutant RAS cell lines. Finally, Arhgef2 is required for mesenchymal transition in a murine mammary model of TGF-induced EMT, suggesting that Arhgef2 may contribute to invasion and metastases in human tumors in vivo.

The identification of Arhgef2 as an essential gene in different tumor types demonstrates that Arhgef2 can mediate cell survival in varied genetic contexts. This idea is supported by the observation that Arhgef2 was not exclusively or inclusively essential in all RAS-mutated cell lines analyzed (Marcotte et al., 2012). This could be explained by the fact that human epithelial tumor cell lines contain diverse genetic aberrations that may confer differential oncogenic dependencies for survival. Such bifurcation of survival dependencies has been shown in pancreatic and lung cell lines, in which the dependence on K-RAS for cell viability is associated with epithelial differentiation state (Singh et al., 2009). In well-differentiated PDAC and lung cell lines, K-RAS expression is required for cell survival. Upon EMT, K-RAS dependency is lost; however, it can be re-gained by MET (Singh et al., 2009). This phenomenon may be

94 explained by the acquisition of additional mutations during the process of EMT that relieve the cell’s dependency on RAS signaling for survival, otherwise known as ‘RAS oncogene addiction’. Since Arhgef2 has been shown to mediate survival and/or proliferation downstream of several pathways including EGF, TNF, taxol, mutant Huntingtin and mutant p53, it is possible that Arhgef2 essentiality may be linked to the relative contribution of one or more of these pathways in addition to mutations in the RAS pathway for cell viability (Kakiashvili et al., 2011, Nie et al., 2012, Varma et al., 2010, Mizuarai et al., 2006). By contrast, our studies in murine fibroblast cells focused on the overexpression of an isolated mutant RAS gene, allowing us to question the dependency of RAS on Arhgef2 expression in a more genetically defined manner. The relative importance of Arhgef2 in cell survival in the context of multiple oncogenic pathways activated in human epithelial cancers, therefore, must be more rigorously tested.

An alternate explanation for the differential survival of Arhgef2-depleted human epithelial cell lines is the efficiency of Arhgef2 knockdown across cell lines used in the shRNA screen. Many of the hairpins used in the study have not been validated for their efficacy and cell line- dependent differences in gene knockdown with identical shRNA sequences has been well documented (Lebbink et al., 2011). It is therefore possible that the significantly essential RAS- mutated cell lines were more sensitive to Arhgef2 knockdown, resulting in the reduction of Arhgef2 transcripts below a threshold required to maintain cell viability. It is unlikely, however, that the reduction in cell survival was the result of off-target effects, since the gene essentiality score (GARP) is the combination of two highest scoring independent shRNAs and we validated the top scoring cell lines with a distinct shRNA sequence that efficiently suppresses Arhgef2 expression levels (Koh et al., 2012).

We also observed a correlation between progressive tumor grade and Arhgef2 expression in human TMAs from lung, colon and pancreatic tumors, malignancies that most frequently harbor RAS mutations (Aguirre et al., 2003, Haigis et al., 2008, Johnson et al., 2001). Although we established that the RAS/MAPK pathway can regulate Arhgef2 expression, the enhanced expression of Arhgef2 observed in late-staged tumorigenesis in vivo may be a result of cooperative oncogenic events involving K-RAS and p53. Gain-of-function mutants of p53 V157F, R175H and R248Q have been shown to transcriptionally upregulate Arhgef2 in NSCLC cell lines and are found in over 50% of late-stage PDAC, NSCLC and colorectal tumors in vivo

(Mizuarai et al., 2006, Maitra et al., 2003, Johnson et al., 1993, Rodrigues et al., 1990). Oncogenic K-RAS and p53 are both required for the malignant conversion of PDAC, suggesting that their co-operative upregulation of Arhgef2 may be one mechanism by which they work together to promote a more malignant phenotype.

Another potential influence of Arhgef2 expression involves the transcriptional induction of Arhgef2 by TGF (Tsapara et al., 2010). Interestingly, approximately half of pancreatic adenocarcinomas undergo homozygous deletion of the TGF signaling component smad4 at late stages of neoplastic progression and Arhgef2 upregulation was found to be smad-dependent (Hezel et al., 2006, Tsapara et al., 2010). However, previous reports have also shown that smad4 is required for TGF-induced EMT in a subset of PDAC and that those tumors that lose smad4 expression retain a well differentiated histopathology (Bardeesy et al., 2006). These data suggest that Arhgef2 upregulation by TGF may co-operate with oncogenic RAS to induce malignant conversion and that Arhgef2 may not contribute to EMT in tumors lacking smad4. However, this idea is challenged by the fact that known effectors of Arhgef2, RhoA and ERK, represent smad- independent pathways that are required for TGF-induced EMT in mammary epithelial cell models and in PDAC models (Bhowmick et al., 2001, Xie et al., 2004, Ellenrieder et al., 2001, Kusama et al., 2001). Moreover, although smad4 is commonly lost, TGF receptors are often overexpressed in PDAC and enhanced TGF signaling correlates with decreased survival (Friess et al., 1993). The contribution of Arhgef2 to the interplay of RAS and TGF signaling pathways in EMT is likely a complex process and whether or not smad signaling is required for Arhgef2- mediated EMT remains to be determined. Ultimately, the cross-talk between Ras, p53 and TGF signaling in the context of pancreatic tumorigenesis and the regulation of Arhgef2 expression suggest that several mechanisms may contribute to the amplification of Arhgef2 expression and attest to the multiple roles of Arhgef2 at different stages of tumor progression (Figure 3.10).

Figure 3.10 Arhgef2 may promote EMT via its cooperative regulation by RASV12, p53 and TGF signaling pathways. Arhgef2 transcripts are induced by oncogenic RAS/MAPK pathway and mutant p53. Increased TGF signaling in the absence of smad4 may result in the enhanced activation of smad-independent pathways, including RAS, ERK1/2 and RhoA. Arhgef2 may (represented as dotted black arrow) mediate TGF-induced RhoA activation and potentiate RASV12/KSR-1/MAPK signaling simultaneously due to elevated expression levels and/or specific activation by both TGF and oncogenic RAS. Increased ERK1/2 and RhoA activity results in increased cell survival, cell migration, gene transcription and ultimately, EMT. Work by Singh et al. showed that RAS-mutated cell lines retaining epithelial characteristics were dependent on RAS for survival, whereas those with mesenchymal properties are insensitive to RAS depletion (Singh et al., 2009). In our study, we showed that Arhgef2 depletion in two mesenchymal-like, or K-RAS independent, cell lines (PANC-1 and A549) induced their reversion to an epithelioid morphology and the re-expression of E-cadherin, whereas Arhgef2 depletion produced little effect on cell lines retaining an epithelial cell morphology (HPAF-II and H727). These data suggest that inhibition of Arhgef2 is able to influence the switch from K- RAS-independency to dependency, thereby re-sensitizing them to K-RAS depletion and

97 subsequent cell death. Thus, the concomitant silencing of Arhgef2 and K-RAS dependent genes may offer novel a mechanism to target K-RAS-independent NSCLC and PDAC tumors.

Although we found that the expression of a distinct RhoA activator could partially compensate for a loss of Arhgef2 expression in TGF-induced EMT, the precise mechanism underlying the requirement of Arhgef2 in this process has not been fully resolved. Both MAPK and RhoA activation are required for TGF-induced EMT in NMuMG cells and our work has shown that Arhgef2 can potentiate both signaling pathways, albeit in context-specific manners (Bhowmick et al., 2001, Xie et al., 2004). Since increased RhoA activity does not fully rescue the mesenchymal phenotype in Arhgef2-depleted cells, it is possible that Arhgef2 impinges on both signaling pathways to promote EMT. Further studies must determine if Arhgef2 mediates TGF- induced ERK1/2 activation, since we have only established a role for Arhgef2 in MAPK pathway activation in the context of oncogenic RAS signaling or PDGF stimulation, and in a KSR-1-dependent manner. Observations that TGF can activate RAS and is dependent on mutated RAS for EMT in some cellular contexts, however, suggests that TGF could indirectly impinge on Arhgef2/KSR-1 signaling in the context of a mutant RAS gene (Hartsough et al., 1996, Frey et al., 1997). Furthermore, both ERK1/2 and RhoA activation have been shown to contribute to EMT in PDAC, where a role for Arhgef2 in ERK activation has been elucidated (Ellenrieder et al., 2001, Kusama et al., 2001). Previous reports have also shown that MEK inhibition induces a partial reversion TGF-induced EMT, as we observed with the ROCK inhibitor Y27632 (Xie et al., 2004). However, we noted a robust prevention of TGF-induced EMT with our most efficient Arhgef shRNA, supporting the notion that Arhgef2 may block several pathways contributing to mesenchymal conversion.

The mechanism by which Arhgef2 mediates the suppression of E-Cadherin expression remains to be resolved. We have shown that Arhgef2 inhibits E-cadherin functionally by preventing its membrane localization, and/or at the molecular level by reducing its gene expression, in a cell type-dependent manner (PDAC/ADC and NMuMG, respectively). It is possible that in mesenchymal cells, depletion of Arhgef2 affects the expression or activation of a mesenchyme- specific transcription factor that suppresses E-cadherin expression, such as snail or slug (snail2). Snail and slug are potent repressors of E-cadherin transcription and inducers of EMT (Batlle et al., 2000, Cano et al., 2000, Nieto et al., 2002). Moreover, snail expression is regulated via the

cooperation of ERK and RhoA pathways (via NFB transcription factors) and in response to oncogenic RAS, all of which are regulated by or impinge on Arhgef2 activity (Barbera et al., 2004, Broders-Bondon et al., 2007). While the minimal ERK binding region of the snail promoter was sufficient to maintain a mesenchymal phenotype in epithelial tumor cells, expression levels of snail did not reach those in cells containing the full promoter, demonstrating that both pathways cooperate to induce snail expression (Barbera et al., 2004). In differentiated epithelial cells, however, Arhgef2 may promote EMT by decreasing the integrity and adhesiveness of tight junctions, an effect that has been previously reported (Benais-Pont et al., 2003, Birukova et al., 2006, Guillemot et al., 2008). This branching of Arhgef2 function depending on differentiation state is supported by the observation that Arhgef2 depletion does not affect E-Cadherin expression in well-differentiated PDAC and ADC cell lines HPAF-II and H727 (Figures 3.7G and 3.7H) as well as NMuMG epithelial cells (Figure 3.9A). Thus, targeting Arhgef2 may prevent both the EMT and the malignant progression of a poorly differentiated tumor by suppressing distinct pathways.

Considering the essential role of Arhgef2 in RAS-driven primary tumorigenesis and EMT, Arhgef2 may be an effective therapeutic target in both early and advanced stages of tumor progression. Given the potential for Arhgef2 to revert RAS-independent tumors back to RAS- dependency by initiating their morphologic de-differentiation, Arhgef2 depletion in conjunction with the inhibition of RAS-essential genes may result in improved therapeutic response in metastatic disease. Ultimately, the development of Arhgef2-directed therapeutics has potential to reduce the malignancy of late-staged PDAC, in which Arhgef2 expression is highest and where current strategies are most ineffective.

Chapter 4

Future Perspectives

4.1 Abstract

We have gained considerable mechanistic insights into the cooperation of Arhgef2 and RAS in tumorigenesis; however, many questions remain unanswered. These questions offer exciting new avenues of research for future students, post-doctoral fellows and/or research associates in our laboratory. While the following suggestions are by no means exhaustive, they are key concepts to address in order to attain a more comprehensive understanding of the role of Arhgef2 in human cancer. First, we must determine the role of Arhgef2 in metastases in vivo, as this is a critical measure of cancer-associated lethality. In addition to PDAC, NSCLC and/or colorectal cancer models should be assessed, since they harbor a high frequency of K-RAS mutations and exhibit elevated Arhgef2 expression with advanced tumor grade. Second, given the transcriptional regulation of Arhgef2 by gain-of-function mutants of p53, it would be of interest to dissect the potential cooperation of these oncogenes in human tumors. A third prospect would be to determine whether Arhgef2 can modify a tumor’s response to established anti-cancer drugs by studying the regulation of Arhgef2 by microtubule-regulating chemotherapeutic agents. Lastly, in order for basic research on the oncogenic function of Arhgef2 to be clinically valuable, we must validate the tractability of Arhgef2 as a therapeutic target.

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4.2 Experimental Procedures

Cell lines and cell culture: OVCAR5, OVCAR8, OVCA420, OVCA429, OVCA432, OVCA433, OCC-1, HOC-1, OVCAR3, CaOV3, CaOV3d3, IOSE80, IOSE29, ES-2, DOV13, A2780, IGROVI, A1847, SK-OV-3IP38, SK-OV-3Bcl2 and SK-OV-3 cell lines were obtained from Gordon Mills and cultured in RPMI 1640 (Life Technologies Inc.) supplemented with 10% FBS (OVCAR5, OVCAR8, DOV13, IOSE80, IOSE29, A1847), McCoy’s 5A Modified Medium (Life Technologies Inc.) supplemented with 10% FBS (OVCA420, OVCA432, OVCAR433, IGROVI), DMEM (Life Technologies Inc.) supplemented with 10% FBS (CaOV3, CaOV3d3, A2780, SK-OV-3, SK-OV-3IP38, SK-OV-3Bcl2) or Alpha Modified Eagle’s Medium (Life Technologies Inc.) supplemented with 10% FBS (OVCA429, OCC1, HOC1, OVCAR3, ES-2). Stable human ARHGEF2 depletion in OVCAR5 cells were generated as described in Chapter 3 and selected with 4g/ml puromycin.

Western blotting: Cells were lysed directly in 2X sample buffer containing 62.5mM Tris-HCl pH 6.8, 2.5% SDS, 10% glycerol, 5% -mercaptoethanol and 0.02% bromophenol blue, boiled and resolved by SDS-PAGE.

Antibodies: Western blotting was performed using monoclonal anti-Arhgef2 antibody 3C5, anti- tubulin (Molecular Probes) and anti-actin (Sigma). Immunohistochemistry studies were performed using anti-Arhgef2 3C5 and anti-p53 (CST, 4937) at 1:500 dilutions.

Immunohistochemistry: Tumor sections derived from normal ovarian tissue and clear cell, mucinous, endometrioid and serous ovarian tumors were prepared and stained as described in Chapter 3. Tumors were given two scores based on intensity of staining and percentage of total tumor stained, with 0 being weak, 1 moderate and 3 strong. Immunoscores were combined to generate scores of 0-6 with 0-2 classified as weak, 3-4 as moderate and 5-6 as high. A total of 4 normal, 22 and 15 clear cell, 17 and 13 mucinous, 26 and 14 endometrioid and 128 and 100 serous tumors were analyzed for Arhgef2 and p53 expression, respectively.

Genome-wide shRNA screen: as described in Marcotte et al., 2012.

Genomic RNA isolation, fragmentation, reverse transcription and amplification in SOC cell lines: mRNA was extracted from OC cell lines using RNeasy plus mini kit (QIAGEN, CN

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74134). mRNA was quantified using QubitRNA broad range and quality control was assessed with bioanalyzer total RNA nano chip.1ug of mRNA was fragmented to an average length of 200bp by incubation 5min at 94°C with 5X fragmentation buffer (Illumina, CN RS-100-0801). Efficiency of the fragmentation was defined on Bioanalyzer RNA Pico Chip. The fragmented mRNA was randomly primed and reverse transcribed using Super Script II cDNA synthesis kit (Invitrogen, CN 18064-014). After second-strand synthesis, the cDNA went through end-repair and ligation reactions according to the Illumina mRNA-Seq Sample Prep Kit protocol. The cDNA library was size-fractioned on a 2% TBE agarose gel. Material in the 350-400bp range was excise and purify (Zymo Research, CN D4001). Half of the eluted cDNA library was used as a template for amplification according mRNA-Seq Sample Prep Kit protocol. The PCR product was purified using the PureLink PCR micro purification kit (invitrogen, catalog no. K310050). The library size (350-400bp) was validated on a Bioanalyzer DNA 1000 Chip and the concentration was estimated using Qubit fluorometer and Quant-iT dsDNA BR Assay Kit (invitrogen, catalog no.Q32850). The library was then used to build clusters on the Illumina flow cell and analysis was done using Illumina Hiseq 2000.

Mapping cDNA fragments and Arhgef2 transcript abundance estimation: Basecalls files were converted to sequences in FASTQ format using BCLToFastq CASAVA 1.8.2. Fragments were mapped to build GRCh37 of the human genome using TopHat 1.4.1 and Bowtie 1.0. Transcript abundance was determined using cufflinks 1.3.0 in Fragment Per Kilobase per Million reads (FPKM). Because alignment was done on a genome version including random chromosomes a correction was applied to the genes present several times. The corrected FPKM values were defined by multiplying the FPKM value of the nonrandom chromosome by the number of times the gene was represented.

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4.3 Future Perspectives

4.3.1 The role of Arhgef2 in metastases

Like the multistep model of primary tumorigenesis, the process of metastasis can be divided into distinct and genetically defined stages (Chiang et al., 2008). The initiation of metastasis involves the invasion of tumor cells through the basement membrane and intravasation through capillary walls into the circulation. Metastatic progression is defined by the survival and extravasian of cancers cells through vessel walls and into distant organs. Finally, in order for metastatic tumors to form, cancer cells must be able to seed and grow at their target sites, a process deemed metastatic virulence (Chiang et al., 2008). EMT promotes the malignant conversion of primary tumor cells and thus is a metastasis initiating event. As such, genes implicated in EMT do not necessarily predict the full metastatic potential of a cancer cell. In order to ascertain the role of Arhgef2 in metastases, therefore, several additional parameters of metastatic progression must be assessed.

We observed that depletion of Arhgef2 resulted in the reversion of the mesenchymal phenotypes associated with metastatic conversion in human lung and pancreatic adenocarcinoma cell lines. In vitro studies should discern whether this results in decreased migration and/or invasion of tumor cells bearing RAS mutations. Following this, an in vivo model of PDAC metastases is required. This can be achieved by allowing primary PDAC xenograft growth to proceed for longer periods of time and assessing whether Arhgef2-depleted tumors have a decreased incidence of evasion from the primary tumor site. Given that Arhgef2 depletion at the time of initiation largely prevents primary tumor growth, however, an inducible Arhgef2 knockdown system could be employed to test the effect of Arhgef2 depletion subsequent to tumor establishment. In this way, we can discern whether silencing Arhgef2 results in tumor regression and/or the prevention of metastases. The potential for acute depletion of Arhgef2 to revert tumor growth and prevent metastases is supported by reports showing that delivery of KSR-1 antisense oligonucleotides in PANC-1 cells prevents primary tumor growth and effects regression of established tumors upon continuous infusion (Xing et al., 2003). Given that Arhgef2 may interfere with both RAS/KSR-1/MAPK and TGF/RhoA and/or TGF/ERK1/2 signaling pathways, the effect of Arhgef2 depletion on tumor regression may be more pronounced.

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To test the role of Arhgef2 in the progression and virulence of metastasis, tail vein injections of PDAC cells stably expressing a hairpin control or Arhgef2 shRNA could be performed. Injecting cancer cells in the tail vein of mice circumvents the steps of invasion-metastasis and directly measures their colonizing potential. This parameter of cancer growth is especially critical since recent studies have shown that cells can metastasize at early stages of tumor development, thereby challenging the traditional multistep progression model of solid tumor growth (Husemann et al., 2008, Podsypanina et al., 2008). In these studies, mouse models with atypical ductal hyperplasia (ADH, a benign stage) in the mammary glands displayed dissemination to the bone marrow (Podsypanina et al., 2008, Husemann et al., 2008). These data argue that metastatic conversion may occur continually throughout the course of primary tumor development, generating a genetically and morphologically diverse spectrum of disseminated cells. In the case of PDAC, the phenomenon of early dissemination would explain its high rate of metastases, refractility to therapeutics and early mortality. Moreover, these studies imply that genes involved in multiple stages of tumorigenic progression would have the most potential as therapeutic targets and thus highlight the importance of elucidating the role of Arhgef2 at later stages of tumorigenic progression.

In addition to ascertaining the biological role of Arhgef2 in malignant progression, we must gain a greater understanding of the biochemical mechanism underlying the contribution of Arhgef2 to EMT and/or PDAC metastases. Arhgef2 can activate Rho GTPases and MAPK signaling, two important promoters of metastatic conversion. Determining the relative importance of these pathways in Arhgef2-mediated EMT and/or malignant progression would provide insight on the metastatic dependencies of PDAC at the molecular level. It is possible that at high expression levels caused by the cooperative upregulation of Arhgef2 by oncogenic RAS and p53, Arhgef2 could efficiently activate both the MAPK and Rho GTPase signaling pathways. In conjunction or alternatively to this idea, the additional TGF and/or mutant p53 signals often seen in advanced stage PDAC may direct Arhgef2 to different substrate pools in the cell. To tease out the Arhgef2- dependent signaling pathways in PDAC cells, TGF and mutant p53-mediated RhoA and ERK1/2 activation must be measured with and without oncogenic RAS and in the presence or absence of Arhgef2 expression. In this way, we will be able to discern whether mutant p53,

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TGF and RAS induce a bifurcation of Arhgef2-mediated signaling events or cooperatively enrich one arm of Arhgef2 signaling.

Lastly, genetic models of RAS-induced tumorigenesis would provide more robust insight into the role of Arhgef2 in RAS tumorigenesis and metastases. A PDAC mouse model has been developed that conditionally expresses oncogenic K-RAS in the pancreas of mice and where 90% of resultant tumors harbor RAS mutations (Hingorani et al., 2003). These mice develop intraepithelial neoplasias at high penetrance and can progress to invasive and metastatic adenocarcinomas. In addition, Tyler Jack’s laboratory has engineered a conditional mutation in the endogenous mouse K-RAS locus that can be activated by ectopic expression of a Cre recombinase (Jackson et al., 2001). AdenoCre infection of mice harboring two copies of the oncogenic allele results in highly efficient induction of lung tumors within 200 days. Mice with one copy of K-RASD12 have a longer survival rate and display tumors of variable stage, spanning mild hyperplasia to overt carcinoma and closely recapitulating human NSCLC. Given the elevated expression of Arhgef2 we observed in late stage NSCLC, this would be a highly relevant model in which to study the role of Arhgef2 in primary lung tumor growth and metastases. Thus, genetic crosses of K-RASD12 PDAC and NSCLC mice with our ARHGEF2 knockout mice would provide excellent model systems to formally test the requirements of ARHGEF2 in K-RAS-mediated tumor induction, growth, metastasis and survival.

4.3.2 The cooperation of Arhgef2 with mutant p53 p53 is a pleiotriopic transcription factor that plays a critical role in preventing tumor cell growth. It lies at the heart of stress response pathways induced by DNA damage, telomere attrition, oncogene activation, hypoxia and aberrant growth signals and functions to restore proper cell function by inducing cell cycle arrest, DNA repair and/or apoptosis (Oren and Rotter, 2010). It is the only gene to surpass the RAS genes in frequency of genetic aberrations in human cancer, with over 50% of all tumors exhibiting loss-of-function or missense mutations in p53 (Hollstein et al., 1991). Moreover, there is mounting evidence that missense p53 mutations not only lose wild-type p53 function but also acquire gain-of-function (GOF) transcriptional and biological activities, thus endowing cancer cells with a double oncogenic hit (Oren and Rotter, 2010).

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ARHGEF2 was identified as a transcriptional target of GOF mutants of p53 (p53V157F, p53R175H and p53R248Q) and promotes the proliferation of NSCLC cells harboring these mutations (Mizuarai et al., 2006). These data suggests that Arhgef2 may cooperate with mutant p53 in cellular transformation and present a powerful new avenue of Arhgef2 research.

Ovarian cancer (OC) encompasses a diverse set of tumor types that exhibit distinct morphological and genetic features. These include clear cell, mucinous, endometrioid and serous ovarian carcinomas (SOC) subtypes, the latter of which account for two-thirds of all ovarian cancers and displays the highest mortality rate (Bernardini et al, 2010). Like PDAC, SOC is often diagnosed at a very late stage when metastases are already present. p53 mutations occur in 60-70% of SOCs, in contrast to the other OC subtypes that exhibit a much lower incidence of p53 mutations and commonly present at an early stage (Havrilesky et al., 2003, Leitao et al., 2004). Although the combined frequency of p53 GOF and null mutations between early and late stages of SOC is comparable, the fraction of missense mutations is significantly higher at advanced stages of tumor progression. Moreover, missense p53 mutations in advanced stage SOC correlate with decreased survival (Bernardini et al., 2010). These studies suggest that gain- of-function mutations in p53 play a driving role in the malignant conversion and associated lethality of SOC.

Given the transcriptional regulation of Arhgef2 by GOF mutants of p53, we sought to determine whether Arhgef2 protein levels were increased in SOC tumors. To that end, we analyzed TMAs derived from human clear cell, mucinous, endometrioid and serous ovarian carcinomas for both Arhgef2 and p53 expression by immunohistostaining (Figure 4.1A). Overexpression of p53 is considered a surrogate for mutant p53 expression, as normal cells express low levels of wild-type p53 and cancer cells are most commonly p53 null or express high levels of the mutant form. Both the staining intensity and percentage of tumor stained were given scores of 0-3 and combined to yield immunoscores ranging from 0-6 for each tumor. Scores of 0-2 were considered weak, 3-4 moderate and 5-6 high. Importantly, we found that Arhgef2 was markedly and specifically upregulated in SOC and stained weakly in all other OC subtypes. Only 5/22 (22.73%) of clear cell, 2/17 (11.76%) of mucinous and 5/26 (19.23%) of endometrioid tumors stained highly for Arhgef2, while 77/128 (60.16%) of SOC tumors displayed high Arhgef2 staining (Figure 4.1B). Moreover, p53 expression exhibited a similar trend, with 2/15 (13.33%)

106 clear cell, 4/13 (30.77%) mucinous and 3/14 (21.43%) endometrioid tumors staining highly for p53 and 62/100 (62%) of SOC tumors showing high levels of p53 staining. These data show a strong correlation between Arhgef2 and p53 expression and suggest that Arhgef2 may functionally contribute to p53-mediated tumor progression in SOC.

Figure 4.1 Arhgef2 is highly expressed in serous ovarian carcinoma. (A) Representative images of tumor microarrays derived from normal ovarian tissue and clear cell, mucinous, endometrioid and serous ovarian carcinomas stained for Arhgef2 expression using a monoclonal antibody against human Arhgef2. Staining is depicted in brown. (B) Distribution of Arhgef2 (left) and p53 (right) immunoscores in OC subtypes represented in (A). Immunoscores were determined based on staining intensity (0-3) and percentage of tumor stained (0-3) to

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generate combined histoscores ranging from 0-6, with 0-2 classified as weak, 3-4 as moderate and 5-6 as strong. n indicates the number of tumors analyzed within each OC subtype. Interestingly, 4 out of the 14 Arhgef2-essential cell lines identified in the genome-wide shRNA screen described in Chapter 3 (Figure 3.4.2A) were of ovarian origin, closely following the 5/14 PDAC cell lines that were essential for Arhgef2. Together, OC and PDAC constituted 64.29% of the cancer types that require Arhgef2 for survival. Moreover, all OC cell lines (4/4) carry mutations in p53 (Table 4.1) with 10 of all 14 Arhgef2-essential cell lines (71.43%) harboring p53 mutations (Samouelian et al., 2004, Redston et al., 1994, O’Connor et al., 1997, Letourneau et al., 2012, Gelfi et al., 1997, Schumacher et al., 1999, Berrozpe et al., 1994, Wasielewski et al., 2006). OV-90 and CFPAC-1, the ovarian and pancreatic cell lines most significantly essential for Arhgef2 (p<0.001 and p=0.003, respectively), carry GOF mutations in both Raf and p53 and K- RAS and p53, respectively (Estep et al., 2007, Samouelian et al., 2004, Kita et al., 1991, Redston et al., 1994). These data suggest that both mutant p53 and RAS require Arhgef2 for survival and that mutation of both oncogenes results in an enhanced dependence on Arhgef2. Moreover, preliminary studies have shown that stable depletion of Arhgef2 in the SOC cell line OVCAR5, harboring wild-type RAS and a mutation in codon 224 of p53 (p53224Q) results in a pronounced decrease in cell viability relative to hairpin control-expressing cells, showing that Arhgef2 essentiality can be validated in at least one model of SOC (Figure 4.2) (O’Connor et al., 1997).

Table 4.1. RAS/MAPK and p53 mutations in ARHGEF2-essential cell lines

Cell line Cancer type *RAS/MAPK status p53 status zGARP p-value OV-90 Ovarian B-RafV600E 1p53S215R -1.49 <0.001 CFPAC-1 Pancreatic K-RASD12 2p35C242R -0.45 0.003 SK-OV-3 Ovarian WT 3p53H179R -0.48 0.006 HCT116 Colorectal K-RASD13 3WT -0.25 0.006 TOV-3133G Ovarian WT 4p53C574T -0.78 0.014 OVCA432 Ovarian WT 5p53C277F -0.89 0.015 Hs578T Breast H-RASV61 6p53V157F -0.42 0.016 Panc 02_03 Pancreatic K-RASD12 DNF -0.7 0.022 PaTu_8988T Pancreatic K-RASD12 7p53C282T -0.31 0.025 IMIMPC-1 Pancreatic K-RASD12 8p53L130V -0.37 0.029 RWP-1 Pancreatic K-RASD12 8p53R175H -0.54 0.032 HRE1 Lung WT DNF -3.05 0.044 SUM1315 Breast WT 9p53C135F -0.05 0.045 1Samouelian et al., 2004, 2Redston et al., 1994, 3O’Connor et al., 1997, 4Letourneau et al., 2012, 5Gelfi et al., 1997, 6Kovach et al., 1991, 7Schumacher et al., 1999, 8Berrozpe et al., 1994, 9Wasielewski et al., 2006 *References stated in Section 3.4.1. DNF = data not found

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Figure 4.2 Arhgef2 is essential for survival in OVCAR5 cells. OVCAR5 cells were stably infected with a hairpin control (shGFP) or hairpin targeting human ARHGEF2 (shGEF) and selected with puromycin for 48h. Representative cell densities are depicted (left) and Arhgef2 protein depletion was confirmed by Western blot (right). Tubulin serves as a protein loading control.

Figure 4.3 Arhgef2 gene expression correlates with essentiality in serous ovarian carcinoma. Arhgef2 gene- essentiality (zGARP) scores obtained from The RNAi Consortium (TRC) across 30 serous ovarian cancer cell lines were plotted against ARHGEF2 transcript levels determined by RNA-Seq analysis.

To discern the potential relationship between mutant p53-mediated Arhgef2 upregulation and Arhgef2 essentiality in SOC cells, we compared the gene essentiality scores (zGARP) with the mRNA expression profiles (FPKM) of ARHGEF2 across 30 SOC cell lines contained within the

109 shRNA screen (Figure 4.3). We found a significant correlation between ARHGEF2 RNA expression and essentiality (R=0.72), suggesting that Arhgef2 transcript upregulation may contribute to increased cell survival in SOC. Together, these data support the hypothesis that ARHGEF2 may be an essential gene in p53-mutated SOC and provide a solid foundation on which to build a future project that could potentially be developed in a parallel manner to our studies investigating Arhgef2 essentiality in K-RAS mutant cell lines and xenograft models. We have analyzed Arhgef2 protein expression in a panel of OC cell lines (Figure 4.4), a cohort of which could be employed to complete further in vitro and in vivo studies to validate the role of Arhgef2 in SOC. Importantly, comparing the effects of OC lines expressing wild-type/null versus GOF mutations in p53 may reveal the potential Arhgef2-dependency of GOF mutants of p53.

Figure 4.4 Arhgef2 expression in human ovarian carcinoma cell lines. Whole cell lysates derived from ovarian carcinoma cell lines were probed for Arhgef2 protein expression by Western blot. Actin serves as a protein loading control.

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4.3.3 The regulation of Arhgef2 by anti-mitotic chemotherapeutic agents

The microtubule cytoskeleton is an effective and validated target for cancer chemotherapeutic drugs. A wide range of structurally distinct molecules can interact with microtubules and interfere with their dynamics, leading to cell cycle arrest and apoptosis (Wilson et al., 1999). Two main classes of anti-mitotic chemotherapeutic drugs (AMCDs) exist: those that bind to  and  tubulin heterodimers and prevent their polymerization (vinca alkaloids), and those that bind polymerized microtubules and stabilize them (taxanes). Nocodazole and taxol (paclitaxel) are prototypical members of each class and are widely used in the treatment of haematological and solid malignancies, respectively (Jordan and Wilson, 2004). Although they have demonstrated potent antitumor activity across many cancer types, they display variable sensitivity in the clinic due to inherent or acquired chemotherapeutic resistance. Thus, the development of agents that can circumvent AMCD resistance is critical to achieve therapeutic efficacy.

There are many mechanisms that may lead to AMCD resistance, including overexpression of drug transporters in the cell, altered drug metabolism, decreased sensitivity to apoptotic stimuli, altered binding of the drug to its target and alterations in microtubule dynamics (Gottesman, 2002). Proteins that regulate microtubule dynamics by interacting with tubulin dimers or polymerized microtubules have been shown to modulate the sensitivity of cells to taxol and/or nocodazole. The most well-studied examples are microtubule-associated proteins (MAPs) such as stathmin and MAP4, which promote microtubule destabilization and stabilization, respectively (Belmont and Mitchison, 1996, Chapin et al., 1995). The downregulation of stathmin sensitizes leukemia cells to taxol and increases their resistance to the vinca alkaloid vinblastine (Iancu et al., 2000 and 2001). In contrast, overexpression of stathmin in lung carcinoma cells sensitizes them to the vinca alkaloid vincristine and has no effect on their sensitivity to taxol (Nishio et al., 2001). Moreover, stathmin inhibited in vitro taxol-induced polymerization of microtubules (Larsson et al., 1999). Conversely, overexpression of MAP4 induces microtubule polymerization and is associated with increased and decreased sensitivity to paclitaxel and vinca alkaloids, respectively (Zhang et al., 1998).

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The net microtubule stabilizing or de-stabilizing effect of microtubule regulating proteins can not necessarily predict its sensitivity to each class of AMCDs, however, since the microtubule stabilizing protein Tau was shown to mediate resistance to paclitaxel in breast cancer (Wagner et al., 2005). Depletion of Tau increased the sensitivity of breast cancer cells to paclitaxel and low Tau expression correlated with enhanced sensitivity to paclitaxel in breast cancer patients (Wagner et al., 2005). Wagner et al. found that the binding of Tau to polymerized microtubules interfered with the binding of paclitaxel, suggesting that reduced interaction of paclitaxel with microtubules results in its decreased efficacy. By contrast, MAP4 and paclitaxel do not interfere with each other’s binding, leading to increased microtubule stability and enhanced paclitaxel sensitivity. To increase this complexity, microtubule targeted drugs can synergize with one another despite having overlapping binding sites on tubulin, suggesting that competitive displacement is not the mechanism underlying binding interference (Martello et al., 2000). These studies show that the contribution of microtubule regulating proteins to AMCD sensitivity is a complex process and must be tested directly to ascertain whether they mediate sensitivity or resistance.

In addition to the miss-expression of microtubule regulating proteins, oncogenic signaling pathways can confer AMCD resistance. The RAS/MAPK pathway is a well-established mediator of AMCD resistance through its regulation of microtubule stability, multidrug-1 resistance (MDR-1) gene induction and survival signaling via the upregulation of the Bcl-2 family of pro- survival proteins (Orr et al., 2005). ERK1/2 is activated in response to microtubule disruption (Shinoharah-Gotoh et al., 1991, Schmid-Alliana et al., 1998) and MEK1/2 inhibition increases taxol sensitivity in breast, ovarian, lung, colorectal and prostate cancer cells (Katayama et al., 2007, Qiu et al., 2005, McDaid et al., 2005, McDaid and Horwitz, 2001, Mhaidat et al., 2009, Zelivianski et al., 2003). Although some studies attribute the enhanced sensitivity of MEK and taxol-treated tumors to the MEK-dependent up- and down-regulation of MDR-1 and Bcl-2, respectively (Katayama et al., 2007, McCubrey et al., 2006), others have found that apoptosis was induced independently of these factors (Bartling et al., 2008).

Arhgef2 is localized to the microtubule array and enhances microtubule stability as evidenced by the presence of increased acetylated beta tubulin and microtubule bundling (Krendel et al., 2002, Schiff and Horwitz, 1980). Importantly, overexpression of Arhgef2 results in increased

112 resistance to the microtubule depolymerising effects of nocodazole and is sequestered to microtubules in response to taxol treatment, demonstrating that Arhgef2 can modulate the effect of these chemotherapeutic drugs (Krendel et al., 2002). Thus, it would be of great interest to discern the effect of Arhgef2 depletion on the sensitivity of cancer cells to both classes of AMCDs. Moreover, given our studies implicating Arhgef2 as a critical mediator of RAS/MAPK signaling in RAS-mutated cancer cells, it is tempting to speculate that Arhgef2 may mediate AMCD resistance by linking microtubule disruption to activation of MAPK signaling. Indeed, our data showing that a cytoplasmically-localized mutant of Arhgef2 can mediate MAPK activation in the absence of oncogenic RAS show that the release of Arhgef2 from microtubules is sufficient to enhance MAPK survival signaling. These observations suggest that Arhgef2 inhibition may sensitize cancer cells to AMCDs and suggest a novel way in which Arhgef2 may serve as a therapeutic target. Given that AMCDs are known to exhibit additive or synergistic effects despite similar mechanisms of action, there is reason to believe that combined inhibition of Arhgef2 with taxol and/or nocodazole may improve therapeutic response. We have obtained preliminary data showing that Arhgef2 is required for nocodazole-induced activation of ERK1/2, thus implicating Arhgef2 in AMCD-mediated activation of MAPK signaling (data not shown). These data greatly strengthen our hypotheses and support the potential of this research to be an exciting future project.

4.3.4 Arhgef2 as a therapeutic target

In our studies we have provided evidence that Arhgef2 may be an effective therapeutic in human cancers harboring RAS mutations. Moreover, the preliminary studies revealed in this chapter suggest that Arhgef2 may also show therapeutic benefit in cancers harboring mutations in p53 and in conjunction with chemotherapeutic agents targeting the microtubule array. In order for these arguments to stand, however, we must ascertain whether Arhgef2 is therapeutically tractable given current technology.

GEFs are not classically considered “druggable,” as they lack the small pockets and grooves required for small molecule interactions (Wells et al., 2007). The elucidation of several GEF- GTPase structures has revealed large and undefined protein-protein interfaces required to

113 facilitate the structural changes of the small GTPase upon binding. This contrasts the small and highly structured catalytic sites of efficiently targeted ATP-binding enzymes. However, the current limitations we face in identifying small molecule inhibitors of GEFs does not mean that it is an impossible feat. In fact, small molecule inhibitors of some GEFs have been identified, supporting the argument that targeting GEFs is achievable. Brefeldin A (BFA), for example, is a drug derived from the fungus Eupenicillium brefeldianum that inhibits Arf GEF by binding to its catalytic domain complexed with Arf-GDP, thereby stabilizing their interaction and preventing subsequent nucleotide exchange (Peyroche et al., 1999, Robineau et al., 2000). This type of inhibition has been termed ‘interfacial inhibition’ and is characterized by the stabilization of protein complexes at or near sites of contact. Moreover, screens have been used to identify peptides that bind to the DH domains of GEFs, exemplified by a peptide targeting the oncogenic GEF TRIO that effectively reduced TRIO-induced tumor formation in xenograft models (Schmidt et al., 2002). Inhibitors of LARG have also been identified, further supporting the notion that GEFs can be targeted by small molecules (Evelyn et al., 2009).

In the case of Arhgef2, one might assume that screens directed at isolating inhibitors of its GEF activity may not be the most desirable since Arhgef2 can activate MAPK signaling independently of its catalytic activity. However, we have demonstrated that Arhgef2 interacts with KSR-1 via its DH domain; therefore, drugs resulting in the inhibition of its GTPase exchange activity may also prevent its interaction with KSR-1 and activation of MAPK signaling. In this way, small molecule inhibitors of the DH domain of Arhgef2 may reduce both MAPK and RhoA signaling and serve as potent chemotherapeutic agents in tumors harboring RAS mutations. Yeast-3-hybrid screens could be performed in which the binding of RhoA to Arhgef2 is assessed in the presence or absence of libraries of small peptides, as was done for TRIO (Schmidt et al., 2002). Positive hits could be subsequently tested for interference of the KSR-1-Arhgef2 interaction to ensure that Arhgef2’s oncogenic GEF-independent functions were also inhibited. These studies would have to be validated in vitro and in vivo for their functional effects and assessed for undesirable off-target effects. Developing an inhibitor for Arhgef2 would undoubtedly be a challenging process; however, with the advancement of high throughput screening methods and improved mechanistic and structural understanding of our desired gene targets, it may be feasible in the years to come.

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While the development of a small molecule inhibitor against Arhgef2 may be attainable, however, it may not reproduce the magnitude of biological effects we observed in our shRNA studies. This possibility is likely given that currently available GEF inhibitors display low potency. Moreover, genetic ablation can differ greatly from pharmacologic inhibition of a target, as evidenced by studies showing that preventing the interaction of RAS and PI3K but not pharmacological inhibition of PI3K is effective in preventing K-RAS-induced lung tumor growth (Gupta et al., 2007).

An alternative strategy to target Arhgef2 in the clinic is through the use of RNA interference (RNAi). Although RNAi has not yet been approved for cancer treatment in humans, it has the potential to be a potent and selective therapeutic modality, as it offers gene-specific targeting. The major barrier to achieving the medicinal potential of RNAi in the form of siRNA lies in its delivery to the cell, as siRNA often display poor stability and non-targeted biodistribution, thereby initiating unwanted immune responses. Recent advances in siRNA delivery methods, however, suggest that siRNA-targeting of Arhgef2 may be a successful means of therapy. A recent publication in the Cullis laboratory showed effective silencing of the androgen receptor (AR) in prostate cancer xenografts via the intravenous (i.v.) injection of lipid nanoparticle (LNP)-encapsulated AR siRNA (Lee et al., 2012). Silencing of AR using shRNA was previously shown to reduce prostate tumor growth and decrease serum prostate specific antigen (PSA) production, thereby implicating AR in prostate tumorigenesis. Importantly, Lee et al. showed that delivery of AR siRNA resulted in efficient endocytosis of the LNP, silencing of AR and decreased serum PSA levels in vivo, demonstrating the feasibility of LNP delivery of siRNA for the efficient silencing of oncogenes in tumors (Lee et al., 2012). Future research investigating the effect of encapsulating Arhgef2 siRNA into LNP systems (Arhgef2LNP) and measuring the initiation and/or regression of pancreatic xenografts following i.v. injection would help unveil the therapeutic potential of Arhgef2.

Current siRNA treatments are largely directed at diseases of the liver, since it is a site of high LNP accumulation. Significantly, the siRNA chemotherapeutic ALN-VSP, containing siRNA directed at kinesin spindle protein (KSP) and vascular endothelial growth factor (VEGF), has shown effect in hepatocellular carcinoma and has recently been promoted to Phase II clinical trials (www.alnylam.com). Given that Arhgef2 was shown to contribute to the malignant

115 progression of HCC, an HCC model of tumorigenesis would serve as an ideal proof-of-concept model to test the efficacy of an Arhgef2LNP (Cheng et al., 2012). A Cullis-Cullis collaboration may therefore put an anti-Arhgef2 chemotherapeutic within clinical reach, and is an essential next step!

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Concluding Remarks

Since the discovery of the RAS genes in the 1980s, we have gained remarkable insight into the molecular complexity of human tumorigenesis. One unifying concept that emerges is that oncogenic signaling is highly context-specific and can evolve throughout the course of tumorigenic progression. While this may seem daunting from a therapeutic perspective, the more we understand these complexities the better we will be at attacking them. New strategies such as combinatorial therapies to target multiple oncogenes in a single tumor and changing therapeutic approaches based on the genetic makeup of a cancer cell – termed personalized medicine – reflect the intellectual advances we have made and are likely to yield improved therapeutic responses in the years to come.

For these reasons, whether or not Arhgef2 is indeed an important determinant of tumorigenic progression does not dictate the value of these studies to the development of effective cancer therapies. The more we understand the subtleties of oncogenic signaling, the closer we will get to achieving curative therapy. From this perspective, I hope to take the knowledge I have gained in the last six years and continue to help the advancement medicinal science in the next phase of my scientific journey.

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Appendix

Appendix 1: Microarray analysis of PANC-1 and H-RASV12-Transformed Fibroblast Cells Harboring Stable Arhgef2 Knockdown

Table I Functional annotation clustering of upregulated genes in Arhgef2-depleted PANC-1 cells

A. Biological Process Count % Enrichment p-value response to organic substance 40 9.80 0.0000 regulation of apoptosis 40 9.80 0.0000 regulation of programmed cell death 40 9.80 0.0000 regulation of cell death 40 9.80 0.0000 response to protein stimulus 13 3.19 0.0000 regulation of cell proliferation 37 9.07 0.0000 biological adhesion 34 8.33 0.0000 anti-apoptosis 16 3.92 0.0001 response to oxidative stress 14 3.43 0.0001 cell adhesion 33 8.09 0.0001 response to inorganic substance 15 3.68 0.0002 response to reactive oxygen species 9 2.21 0.0002 negative regulation of apoptosis 20 4.90 0.0003 negative regulation of programmed cell death 20 4.90 0.0004 negative regulation of cell death 20 4.90 0.0004 negative regulation of cell proliferation 20 4.90 0.0004 regulation of smooth muscle cell proliferation 7 1.72 0.0005 regulation of locomotion 13 3.19 0.0011 cell migration 16 3.92 0.0012 response to mechanical stimulus 7 1.72 0.0014 cell motion 22 5.39 0.0019 cellular response to oxidative stress 6 1.47 0.0024 protein localization at cell surface 3 0.74 0.0028 negative regulation of transcription factor activity 6 1.47 0.0029 response to hormone stimulus 18 4.41 0.0031 cell motility 16 3.92 0.0033 localization of cell 16 3.92 0.0033 response to steroid hormone stimulus 12 2.94 0.0035 positive regulation of chemotaxis 5 1.23 0.0035 regulation of cell migration 11 2.70 0.0042 positive regulation of smooth muscle cell proliferation 5 1.23 0.0045 regulation of chemotaxis 5 1.23 0.0045 response to unfolded protein 7 1.72 0.0046

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negative regulation of DNA binding 6 1.47 0.0051 positive regulation of locomotion 8 1.96 0.0058 positive regulation of behavior 5 1.23 0.0063 cellular homeostasis 20 4.90 0.0072 response to hydrogen peroxide 6 1.47 0.0076 negative regulation of biosynthetic process 23 5.64 0.0079 extracellular matrix organization 8 1.96 0.0079 blood vessel development 13 3.19 0.0080 negative regulation of cell migration 6 1.47 0.0081 leukocyte migration 6 1.47 0.0081 response to endogenous stimulus 18 4.42 0.0082 regulation of leukocyte migration 4 0.98 0.0091 negative regulation of macromolecule biosynthetic process 22 5.39 0.0093 sequestering of metal ion 3 0.74 0.0094 negative regulation of binding 6 1.47 0.0094 vasculature development 13 3.19 0.0096 extracellular structure organization 10 2.45 0.0099

B. Molecular Function Count % Enrichment p-value kinase binding 12 2.94 0.0025 antioxidant activity 6 1.47 0.0040 oxidoreductase activity, acting on peroxide as acceptor 5 1.23 0.0056 peroxidase activity 5 1.23 0.0056 collagen binding 5 1.23 0.0085

C. KEGG Pathway Count % Enrichment p-value Focal adhesion 16 3.92 0.0003 ECM-receptor interaction 8 1.96 0.0077 NOD-like receptor signaling pathway 6 1.47 0.0264 Bladder cancer 5 1.23 0.0268 Epithelial cell signaling in Helicobacter pylori infection 6 1.47 0.0374 Leukocyte transendothelial migration 8 1.96 0.0418

Table II Upregulated gene lists in Arhgef2-depleted PANC-1 cells by functional annotation

A. Regulation of Apoptosis (40) (Biological Process)

ENTREZ Gene ID Gene Name

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694 B-cell translocation gene 1, anti-proliferative 4671 NLR family, apoptosis inhibitory protein 6502 S-phase kinase-associated protein 2 (p45) 51282 SCAN domain containing 1 55437 STE20-related kinase adaptor beta 51499 TP53 regulated inhibitor of apoptosis 1 301 annexin A1 317 apoptotic peptidase activating factor 1 999 cadherin 1, type 1, E-cadherin (epithelial) 837 caspase 4, apoptosis-related cysteine peptidase 6347 chemokine (C-C motif) ligand 2 162989 death effector domain containing 2 1611 death-associated protein excision repair cross-complementing rodent repair deficiency, 2073 group 5 51083 galanin prepropeptide 3336 heat shock 10kDa protein 1 (chaperonin 10) 3304, 3303 heat shock 70kDa protein 1A; heat shock 70kDa protein 1B 7184 heat shock protein 90kDa beta (Grp94), member 1 3162 heme oxygenase (decycling) 1 348 hypothetical LOC100129500; apolipoprotein E inhibitor of DNA binding 3, dominant negative helix-loop-helix 3399 protein 3482 insulin-like growth factor 2 receptor 9445 integral membrane protein 2B 3570 interleukin 6 receptor 4318 matrix metallopeptidase 9 5601 mitogen-activated protein kinase 9 4487 msh homeobox 1 27018 nerve growth factor receptor (TNFRSF16) associated protein 1 26471 nuclear protein, transcriptional regulator, 1 7001 peroxiredoxin 2 10935 peroxiredoxin 3 5051 platelet-activating factor acetylhydrolase 2 5621 prion protein 5578 protein kinase C, alpha 9616 ring finger protein 7 6609 sphingomyelin phosphodiesterase 1 10628 thioredoxin interacting protein 7057 thrombospondin 1 7009 transmembrane BAX inhibitor motif containing 6 7428 von Hippel-Lindau tumor suppressor

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B. Biological Adhesion (32) (Biological Process)

ENTREZ Gene ID Gene Name 83692 CD99 molecule-like 2 8857 Fc fragment of IgG binding protein 9289 G protein-coupled receptor 56 5754 PTK7 protein tyrosine kinase 7 999 cadherin 1, type 1, E-cadherin (epithelial) 1000 cadherin 2, type 1, N-cadherin (neuronal) 1001 cadherin 3, type 1, P-cadherin (placental) 56265 carboxypeptidase X (M14 family), member 1 6347 chemokine (C-C motif) ligand 2 9076 claudin 1 9071 claudin 10 9069 claudin 12 1301 collagen, type XI, alpha 1 1829 desmoglein 2 285761 discoidin, CUB and LCCL domain containing 1 131566 discoidin, CUB and LCCL domain containing 2 10979 fermitin family homolog 2 (Drosophila) 3673 integrin, alpha 2 (CD49B, alpha 2 subunit of VLA-2 receptor) 3678 integrin, alpha 5 (fibronectin receptor, alpha polypeptide) 3688 integrin, beta 1 (fibronectin receptor, beta polypeptide) 9235 interleukin 32 3918 laminin, gamma 2 10446 leucine rich repeat neuronal 2 9404 leupaxin 4478 moesin 29780 parvin, beta 5796 protein tyrosine phosphatase, receptor type, K 10076 protein tyrosine phosphatase, receptor type, U 8082 sarcospan (Kras oncogene-associated gene) 113675 serine dehydratase-like 140885 signal-regulatory protein alpha 6695 sparc/osteonectin, cwcv and kazal-like domains proteoglycan 1 7057 thrombospondin 1 7045 transforming growth factor, beta-induced, 68kDa

C. Cell Motion (22) (Biological Process)

ENTREZ Gene ID Gene Name 694 B-cell translocation gene 1, anti-proliferative 2048 EPH receptor B2

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301 annexin A1 1000 cadherin 2, type 1, N-cadherin (neuronal) 800 caldesmon 1 6347 chemokine (C-C motif) ligand 2 1839 heparin-binding EGF-like growth factor inhibitor of DNA binding 1, dominant negative helix-loop-helix 3397 protein 3673 integrin, alpha 2 (CD49B, alpha 2 subunit of VLA-2 receptor) 3678 integrin, alpha 5 (fibronectin receptor, alpha polypeptide) 3688 integrin, beta 1 (fibronectin receptor, beta polypeptide) 3570 interleukin 6 receptor 3576 interleukin 8 4478 moesin 5420 podocalyxin-like 5578 protein kinase C, alpha 5796 protein tyrosine phosphatase, receptor type, K 6695 sparc/osteonectin, cwcv and kazal-like domains proteoglycan 1 7057 thrombospondin 1 7171 tropomyosin 4 7424 vascular endothelial growth factor C 7428 von Hippel-Lindau tumor suppressor

D. Cell Junction (24) (Cellular Component)

ENTREZ Gene ID Gene Name 83692 CD99 molecule-like 2 153562 MARVEL domain containing 2 999 cadherin 1, type 1, E-cadherin (epithelial) 1000 cadherin 2, type 1, N-cadherin (neuronal) 1001 cadherin 3, type 1, P-cadherin (placental) 9076 claudin 1 9071 claudin 10 9069 claudin 12 1829 desmoglein 2 57669 erythrocyte membrane protein band 4.1 like 5 10979 fermitin family homolog 2 2560 gamma-aminobutyric acid (GABA) A receptor, beta 1 3673 integrin, alpha 2 (CD49B, alpha 2 subunit of VLA-2 receptor) 3678 integrin, alpha 5 (fibronectin receptor, alpha polypeptide) 3688 integrin, beta 1 (fibronectin receptor, beta polypeptide) 5058 p21 protein (Cdc42/Rac)-activated kinase 1 24145 pannexin 1

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29780 parvin, beta 23362 pleckstrin and Sec7 domain containing 3 5796 protein tyrosine phosphatase, receptor type, K 10076 protein tyrosine phosphatase, receptor type, U 8082 sarcospan (Kras oncogene-associated gene) 26872 six transmembrane epithelial antigen of the prostate 1 127262 tumor protein p63 regulated 1-like

E. Focal Adhesion (16) (KEGG Pathway)

ENTREZ Gene ID Gene Name 1277 collagen, type I, alpha 1 1290 collagen, type V, alpha 2 1301 collagen, type XI, alpha 1 2316 filamin A, alpha (actin binding protein 280) 3673 integrin, alpha 2 (CD49B, alpha 2 subunit of VLA-2 receptor) 3678 integrin, alpha 5 (fibronectin receptor, alpha polypeptide) 3688 integrin, beta 1 (fibronectin receptor, beta polypeptide) 3918 laminin, gamma 2 5601 mitogen-activated protein kinase 9 10398 myosin, light chain 9, regulatory 5058 p21 protein (Cdc42/Rac)-activated kinase 1 56924 p21 protein (Cdc42/Rac)-activated kinase 6 29780 parvin, beta 5578 protein kinase C, alpha 7057 thrombospondin 1 7424 vascular endothelial growth factor C

F. ECM-Receptor Interaction (8) (KEGG Pathway)

ENTREZ Gene ID Gene Name 1277 collagen, type I, alpha 1 1290 collagen, type V, alpha 2 1301 collagen, type XI, alpha 1 3673 integrin, alpha 2 (CD49B, alpha 2 subunit of VLA-2 receptor) 3678 integrin, alpha 5 (fibronectin receptor, alpha polypeptide) 3688 integrin, beta 1 (fibronectin receptor, beta polypeptide) 3918 laminin, gamma 2 7057 thrombospondin 1

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Table III Functional annotation clustering of downregulated genes in Arhgef2-depleted PANC-1 cells

A. Biological Process Count % Enrichment p-value positive regulation of organelle organization 10 2.88 0.0000 mesenchymal cell development 8 2.31 0.0000 mesenchymal cell differentiation 8 2.31 0.0000 mesenchyme development 8 2.31 0.0000 regulation of organelle organization 15 4.32 0.0000 M phase 18 5.19 0.0001 positive regulation of cytoskeleton organization 7 2.02 0.0002 regulation of transcription from RNA polymerase II promoter 29 8.36 0.0002 cell cycle phase 20 5.76 0.0002 negative regulation of cellular biosynthetic process 24 6.92 0.0003 negative regulation of biosynthetic process 24 6.92 0.0004 regulation of phosphorylation 21 6.05 0.0004 cell cycle 29 8.36 0.0005 regulation of nuclear division 7 2.02 0.0005 regulation of mitosis 7 2.02 0.0005 regulation of anti-apoptosis 6 1.73 0.0006 regulation of phosphate metabolic process 21 6.05 0.0006 regulation of phosphorus metabolic process 21 6.05 0.0006 cell cycle process 23 6.63 0.0007 regulation of cytoskeleton organization 10 2.88 0.0009 regulation of kinase activity 17 4.90 0.0009 organelle fission 13 3.75 0.0011 negative regulation of macromolecule biosynthetic process 22 6.34 0.0011 cell proliferation 19 5.48 0.0011 mitotic cell cycle 17 4.90 0.0013 regulation of transferase activity 17 4.90 0.0014 negative regulation of nitrogen compound metabolic process 21 6.05 0.0014 establishment of organelle localization 7 2.02 0.0016 regulation of protein kinase activity 16 4.61 0.0018 blood vessel morphogenesis 12 3.46 0.0018 positive regulation of cellular component organization 11 3.17 0.0019 negative regulation of transcription 19 5.48 0.0020 negative regulation of macromolecule metabolic process 26 7.49 0.0021 chromosome localization 4 1.15 0.0024

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establishment of chromosome localization 4 1.15 0.0024 negative regulation of gene expression 20 5.76 0.0024 negative regulation of transcription, DNA-dependent 16 4.61 0.0024 mitosis 12 3.45 0.0025 nuclear division 12 3.46 0.0025 regulation of endothelial cell proliferation 5 1.45 0.0026 negative regulation of RNA metabolic process 16 4.61 0.0028 cytoskeleton organization 18 5.19 0.0028 M phase of mitotic cell cycle 12 3.46 0.0029 negative regulation of nucleobase, nucleoside, nucleotide and nucleic acid metabolic process 20 5.76 0.0029 neural crest cell development 5 1.44 0.0030 neural crest cell differentiation 5 1.44 0.0030 regulation of transcription, DNA-dependent 49 14.12 0.0033 regulation of apoptosis 27 7.78 0.0035 positive regulation of protein complex assembly 5 1.44 0.0037 ectoderm development 11 3.17 0.0037 regulation of programmed cell death 27 7.78 0.0040 epithelial to mesenchymal transition 4 1.15 0.0041 regulation of cell death 27 7.78 0.0042 regulation of cellular component size 13 3.75 0.0043 embryonic organ development 10 2.88 0.0045 skeletal system morphogenesis 8 2.31 0.0045 regulation of cellular component biogenesis 9 2.59 0.0046 negative regulation of cellular component organization 9 2.59 0.0046 regulation of cell cycle process 8 2.31 0.0050 regulation of RNA metabolic process 49 14.12 0.0051 regulation of cell proliferation 26 7.49 0.0052 blood vessel development 12 3.45 0.0056 muscle organ development 11 3.17 0.0056 positive regulation of molecular function 21 6.05 0.0057 angiogenesis 9 2.59 0.0059 skeletal system development 14 4.03 0.0060 regulation of protein complex assembly 7 2.02 0.0062 cellular macromolecular complex subunit organization 15 4.32 0.0062 vasculature development 12 3.46 0.0067 organelle localization 7 2.02 0.0069 epidermis development 10 2.88 0.0069 regulation of mitotic cell cycle 9 2.59 0.0070 transcription, DNA-dependent 13 3.75 0.0077 regulation of protein polymerization 6 1.73 0.0081

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regulation of cell cycle 14 4.04 0.0082 RNA biosynthetic process 13 3.75 0.0086 regulation of locomotion 10 2.88 0.0090

B. Cellular Component Count % Enrichment p-value chromosome, centromeric region 10 2.88 0.0002 kinetochore 8 2.31 0.0003 condensed chromosome kinetochore 7 2.02 0.0004 chromosome 19 5.48 0.0008 cytoskeleton 40 11.53 0.0008 condensed chromosome, centromeric region 7 2.02 0.0008 non-membrane-bounded organelle 64 18.44 0.0010 intracellular non-membrane-bounded organelle 64 18.44 0.0010 chromosomal part 16 4.61 0.0023 spindle pole 5 1.44 0.0024 spindle 9 2.59 0.0034 condensed chromosome 8 2.31 0.0061

C. Molecular Function Count % Enrichment p-value transcription cofactor activity 20 5.76 0.0000 transcription factor binding 24 6.92 0.0001 transcription regulator activity 49 14.12 0.0001 transcription factor activity 35 10.09 0.0002 enzyme binding 23 6.63 0.0002 transcription activator activity 17 4.90 0.0033 GTPase binding 8 2.31 0.0036 GTP-Rho binding 3 0.86 0.0047 Rho GTPase binding 5 1.44 0.0048 protein serine/threonine kinase activity 17 4.90 0.0052 transcription coactivator activity 11 3.17 0.0056 Ras GTPase binding 7 2.02 0.0061 protein kinase activity 21 6.05 0.0070 protein tyrosine kinase activator activity 3 0.86 0.0085

D. KEGG Pathway Count % Enrichment p-value MAPK signaling pathway 12 3.46 0.0054 Oocyte meiosis 7 2.02 0.0109 Focal adhesion 9 2.59 0.0209

Table IV Downregulated gene lists in Arhgef2-depleted PANC-1 cells by functional annotation

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A. Mesenchymal Cell Development/Differentiation (8) (Biological Process)

ENTREZ Gene ID Gene Name

6275 S100 calcium binding protein A4 6662 SRY (sex determining region Y)-box 9 6899 T-box 1 1906 endothelin 1 8320 eomesodermin homolog 3911 laminin, alpha 5 3084 neuregulin 1 nuclear factor of activated T-cells, cytoplasmic, calcineurin- 4772 dependent 1

B. Anti-Apoptosis (6) (Biological Process)

ENTREZ Gene ID Gene Name

581 BCL2-associated X protein 133 adrenomedullin 857 caveolin 1 1843 dual specificity phosphatase 1 6242 rhotekin 23411 sirtuin (silent mating type information regulation 2 homolog) 1

C. Cell Migration (9) (Biological Process)

ENTREZ Gene ID Gene Name Cbp/p300-interacting transactivator, with Glu/Asp-rich 10370 carboxy-terminal domain, 2 7070 Thy-1 cell surface antigen 2152 coagulation factor III 1906 endothelin 1 3486 insulin-like growth factor binding protein 3 3911 laminin, alpha 5 5594 mitogen-activated protein kinase 1 5155 platelet-derived growth factor beta polypeptide 5879 ras-related C3 botulinum toxin substrate 1 (Rac1)

D. Cytoskeleton (4) (Cellular Component)

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ENTREZ Gene ID Gene Name

9590 A kinase (PRKA) anchor protein 12 148170 CDC42 effector protein (Rho GTPase binding) 5 10160 FERM, RhoGEF (ARHGEF) and pleckstrin domain protein 1 3005 H1 histone family, member 0 9181 Rho/Rac guanine nucleotide exchange factor (GEF) 2 11078 TRIO and F-actin binding protein 60312 actin filament associated protein 1 83543 allograft inflammatory factor 1-like 23299 bicaudal D homolog 2 274 bridging integrator 1 55450 calcium/calmodulin-dependent protein kinase II inhibitor 1 801 calmodulin 3 (phosphorylase kinase, delta) 1062 centromere protein E, 312kDa 1063 centromere protein F, 350/400ka (mitosin) 22995 centrosomal protein 152kDa 1808 dihydropyrimidinase-like 2 9787 discs, large (Drosophila) homolog-associated protein 5 2037 erythrocyte membrane protein band 4.1-like 2 79187 fibronectin type III and SPRY domain containing 1 2318 filamin C, gamma 284085 hypothetical protein FLJ40504 149501 keratin 8 pseudogene 9; similar to keratin 8 144501 keratin 80 55329 meiosis-specific nuclear structural 1 5594 mitogen-activated protein kinase 1 4644 myosin VA (heavy chain 12, myoxin) 25924 myosin VIIA and Rab interacting protein 140465 myosin, light chain 6B, alkali, smooth muscle and non-muscle 84276 nicolin 1 54820 nudE nuclear distribution gene E homolog 1 4957 outer dense fiber of sperm tails 2 5062 p21 protein (Cdc42/Rac)-activated kinase 2 5347 polo-like kinase 1 (Drosophila) protein phosphatase 2 (formerly 2A), catalytic subunit, beta 5516 isoform 5925 retinoblastoma 1 10174 sorbin and SH3 domain containing 3 3925 stathmin 1 11013 thymosin beta 15a 7138 troponin T type 1 347733 tubulin, beta 2B

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E. Focal Adhesion (9) (KEGG Pathway)

ENTREZ Gene ID Gene Name

857 caveolin 1 858 caveolin 2 2318 filamin C, gamma 3911 laminin, alpha 5 10319 laminin, gamma 3 5594 mitogen-activated protein kinase 1 5062 p21 protein (Cdc42/Rac)-activated kinase 2 5155 platelet-derived growth factor beta polypeptide 5879 ras-related C3 botulinum toxin substrate 1 (Rac1)

Table V Functional annotation clustering of Significance Analysis of Microarray (SAM) genes downregulated in Arhgef2-depleted NIH 3T3-H-RasV12 cells % A. Biological Process Count Enrichment p-value positive regulation of ERK1 and ERK2 cascade 3 2.46 0.0013 regulation of ERK1 and ERK2 cascade 3 2.46 0.0020 response to wounding 9 7.38 0.0026 inflammatory response 7 5.74 0.0044 taxis 5 4.10 0.0066 chemotaxis 5 4.10 0.0066 epithelial cell differentiation 5 4.10 0.0101 transmembrane receptor protein tyrosine kinase signaling pathway 6 4.92 0.0102 regulation of morphogenesis of a branching structure 3 2.46 0.0126 mammary gland bud formation 2 1.64 0.0136 branch elongation involved in salivary gland morphogenesis 2 1.64 0.0136 fibroblast growth factor receptor signaling pathway 3 2.46 0.0178 salivary gland development 3 2.46 0.0189 mammary gland bud morphogenesis 2 1.64 0.0204 behavior 8 6.56 0.0210 response to organic substance 9 7.38 0.0222 positive regulation of endocytosis 3 2.46 0.0226 lacrimal gland development 2 1.64 0.0271 exocrine system development 3 2.46 0.0320 defense response 8 6.56 0.0338 response to endogenous stimulus 5 4.10 0.0375

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epithelium development 6 4.92 0.0383 enzyme linked receptor protein signaling pathway 6 4.92 0.0393 lung development 4 3.28 0.0405 positive regulation of MAPKKK cascade 3 2.46 0.0411 electron transport chain 4 3.28 0.0415 respiratory tube development 4 3.28 0.0424 cell-matrix adhesion 3 2.46 0.0459 gland development 5 4.10 0.0462 prostate glandular acinus morphogenesis 2 1.64 0.0469 prostate epithelial cord arborization in prostate glandular acinus morph. 2 1.64 0.0469 regulation of branching involved in prostate gland morphogenesis 2 1.64 0.0469 epithelial cell proliferation involved in salivary gland morphogenesis 2 1.64 0.0469 regulation of endocytosis 3 2.46 0.0493

B. Cellular Component Count % Enrichment p-value adherens junction 6 4.92 0.0007 anchoring junction 6 4.92 0.0014 cell junction 10 8.20 0.0042 focal adhesion 4 3.28 0.0066 cell-substrate adherens junction 4 3.28 0.0080 cell-substrate junction 4 3.28 0.0099 actin cytoskeleton 6 4.92 0.0122 muscle thin filament tropomyosin 2 1.64 0.0134 extracellular region part 12 9.84 0.0142 contractile fiber part 4 3.28 0.0201 contractile fiber 4 3.28 0.0260 microsome 5 4.10 0.0307 vesicular fraction 5 4.10 0.0341 extrinsic to membrane 8 6.56 0.0391 organelle membrane 11 9.02 0.0441 striated muscle thin filament 2 1.64 0.0461 cell fraction 9 7.38 0.0464 mitochondrial inner membrane 6 4.92 0.0489

C. Molecular Function Count % Enrichment p-value pattern binding 6 4.92 0.0015 polysaccharide binding 6 4.92 0.0015 chemokine activity 4 3.28 0.0019 chemokine receptor binding 4 3.28 0.0021

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glycosaminoglycan binding 5 4.10 0.0066 growth factor binding 4 3.28 0.0117 heparin binding 4 3.28 0.0171 carbohydrate binding 7 5.74 0.0177 cytokine activity 5 4.10 0.0304

D. KEGG Pathway Count % Enrichment p-value Metabolism of xenobiotics by cytochrome P450 5 4.10 0.0023 Chemokine signaling pathway 7 5.74 0.0042 Tight junction 6 4.92 0.0056 Glutathione metabolism 4 3.28 0.0096 Drug metabolism 4 3.28 0.0257

Table VI Downregulated gene lists in Arhgef2-depleted NIH 3T3-H-RASV12 cells by functional annotation

A. Response to Wounding (9) (Biological Process)

ENTREZ Gene ID Gene Name 20296 chemokine (C-C motif) ligand 2 20306 chemokine (C-C motif) ligand 7 14825 chemokine (C-X-C motif) ligand 1 nuclear factor of kappa light polypeptide gene enhancer in B- 80859 cells inh z 100044702 similar to LPS-induced CXC chemokine; chemokine ligand 5 20848 signal transducer and activator of transcription 3 solute carrier family 1 (glial high affinity glutamate transporter), 20512 member 3 21859 tissue inhibitor of metalloproteinase 3 21898 toll-like receptor 4

B. Epithelial Cell Differentiation (5) (Biological Process)

ENTREZ Gene ID Gene Name 22433 X-box binding protein 1 14165 fibroblast growth factor 10 14183 fibroblast growth factor receptor 2 16664 keratin 14 21804 transforming growth factor beta 1 induced transcript 1

C. Fibroblast Growth Factor Receptor Signaling Pathway (3) (Biological Process)

131

ENTREZ Gene ID Gene Name 14219 connective tissue growth factor 14165 fibroblast growth factor 10 14183 fibroblast growth factor receptor 2

D. Cell-Matrix Adhesion (3) (Biological Process)

ENTREZ Gene ID Gene Name 14219 connective tissue growth factor 16419 integrin beta 5 19261 signal-regulatory protein alpha

E. Adherens Junctions (5) (Cellular Component)

ENTREZ Gene ID Gene Name 65970 LIM domain and actin binding 1 109711 actinin, alpha 1 17356 mixed-lineage leukemia; translocated to, 4 19294 poliovirus receptor-related 2 21753 testis derived transcript 21804 transforming growth factor beta 1 induced transcript 1

F. Cell Junction (10) (Cellular Component)

ENTREZ Gene ID Gene Name 65970 LIM domain and actin binding 1 109711 actinin, alpha 1 13823 erythrocyte protein band 4.1-like 3 17356 mixed-lineage leukemia; translocated to, 4 93737 par-6 partitioning defective 6 homolog gamma 19294 poliovirus receptor-related 2 52398 septin 11 21753 testis derived transcript 60409 trafficking protein particle complex 4 21804 transforming growth factor beta 1 induced transcript 1

G. Tight Junction (6) (KEGG Pathway)

ENTREZ Gene ID Gene Name 109711 actinin, alpha 1 13823 erythrocyte protein band 4.1-like 3

132

17356 mixed-lineage leukemia; translocated to, 4 98932 myosin, light polypeptide 9, regulatory 93737 par-6 partitioning defective 6 homolog gamma 23797 thymoma viral proto-oncogene 3

Tables I-IV. Microarray analysis of Arhgef2-depleted PANC-1 cells. Table I: Gene ontology analysis of the 399 most upregulated genes in PANC-1shGFP vs PANC-1shGEF cells (greater than DF log2 of 0.6 or 1.52 fold-change) showing gene classification based on biological process (Table IA), cellular component (Table IB), molecular function (Table IC) and KEGG pathway (Table ID). Table II: upregulated genes of interest are listed by functional annotation category in Tables IIA-F. Table III: The 416 most downregulated genes in PANC-1shGFP vs PANC- 1shGEF cells (less than DF log2 of -0.6 or -1.52 fold-change) were classified as in Table I in Tables IIIA-D. Table IV: downregulated genes of interest are listed by functional annotation in Tables IVA-E. Count reveals number of genes perturbed within each annotated group, % enrichment denotes the fraction of genes associated with its respective group that are enriched in the dataset. P-values of less than 0.01 or 0.05 are shown.

Tables V and VI. Microarray analysis of Arhgef2-depleted NIH 3T3-H-RASV12 cells. Table V: Gene ontology analysis of 170 significantly downregulated genes in Arhgef2-depleted NIH 3T3-H-RASV12 cells as measured by SAM analysis. Gene enrichment according to biological process, cellular component, molecular function and KEGG pathway are shown in Tables VA-D, respectively. Count, % enrichment and p-values are as stated for Tables I-IV. Table VI: genes of interest are listed by functional annotation in Tables VIA-G.

133

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