Novel Interactions Mediated by the Adaptor Protein Dok-4 Regulate

Novel interactions mediated by the adaptor protein Dok-4

regulate growth factor signaling and could influence the

response to acute kidney injury

Victoria Roodman

Department of Experimental Medicine

McGill University

Montreal, Quebec, Canada

April 2018

A thesis submitted to McGill University in partial fulfillment of the requirements of the degree

of Master of Science.

ABSTRACT Ischemia-reperfusion injury (IRI) causes acute kidney injury, particularly inducing cell death in proximal tubule epithelial cells (TECs). Following injury, surviving TECs regenerate to repair injured tubules, a process which in animal models, is enhanced by endogenous and exogenous growth factors (GFs), such as epidermal growth factor. Unfortunately, GF-based intervention has been disappointing in human trials, suggesting that negative regulatory mechanisms must be overcome to maximize the benefit of these interventions.

We hypothesize that Dok-4, a conserved member of the Dok family of inhibitory adapter proteins, which is highly expressed in epithelial tissues yet remains virtually uncharacterized, mediates anti-mitogenic signaling in renal tubules following IRI by recruiting β2-chimaerin, a negative regulator of the Rho GTPase Rac1.

We have found that Dok-4 and β2-chimaerin both to be upregulated in the injured kidney following ischemia-reperfusion. We have found that Dok-4 interacts with β2-chimaerin and that structurally, the Dok-4/β2-chimaerin interaction occurs directly through the Dok-4 phosphotyrosine-binding (PTB) domain and a phosphorylated tyrosine residue at position 153 (Y153) of β2-chimaerin, and that the interaction is potentiated by tyrosine kinases. We have identified the homologous α2-chimaerin, as containing the same PTB binding motif (NPIY143), and confirmed that, as with β2-chimaerin, it interacts with the Dok-4 PTB domain upon phosphorylation. We also found that the interaction with Dok-4 is greatly enhanced by the active (open) conformation of β2-chimaerin.

Functionally, we have found that Dok-4 and β2-chimaerin cooperate to inhibit Rac1 activity at the cell membrane and that this inhibitory effect requires membrane localization mediated by the Dok-4 pleckstrin homology domain. From these results, we propose a model whereby Dok-4 acts to recruit β2-chimaerin to the membrane, where it is stabilized in its active form and is able to attenuate Rac1 signaling and presumably, downstream proliferative signaling events.

IRI is associated with the progression to end-stage renal disease and negative patient

2 outcome. Understanding the role of Dok-4 in the negative regulation of mitogenic signaling in injured renal tubules will help develop treatments aimed at limiting post-ischemic renal injury and improve patient outcome. Such interventions might include inhibition of Dok-4 expression or interference with the recruitment or activation of β2-chimaerin.

RÉSUMÉ

Une insulte d'ischémie-reperfusion (IRI) provoque une lésion rénale aiguë, en particulier induisant la mort cellulaire dans les cellules épithéliales tubulaires (CET) proximales. Après l’IRI, les CET survivants se régénèrent pour réparer les tubules endommagés, un processus qui, chez les modèles animaux, est renforcé par des facteurs de croissance (FC) endogènes et exogènes, tels que le facteur de croissance épidermique. Malheureusement, l'intervention basée sur les FC a été décevante dans les essais humains, ce qui suggère que des mécanismes de régulation négatifs doivent être surmontés pour maximiser les bénéfices de ces interventions.

Nous émettons l'hypothèse que Dok-4, un membre conservé de la famille Dok des protéines adaptatrices inhibitrices, qui est fortement exprimé dans les tissus épithéliaux mais reste pratiquement non caractérisé, médie la signalisation anti-mitogène dans les tubules rénaux à la suite d’IRI en recrutant β2-chimérine, un régulateur négatif de la Rho GTPase Rac1.

Nous avons trouvé que Dok-4, β2-chimérine et α2-chimérine sont toutes régulées positivement dans le rein après ischémie-reperfusion. Nous avons trouvé que Dok-4 interagit avec la β2-chimérinee et que structurellement, l'interaction Dok-4 / β2-chimérine se produit directement à travers le domaine de liaison à la phosphotyrosine (DLP) Dok-4 et un résidu tyrosine phosphorylé à la position 153 (Y153) de β2-chimérine, et que l'interaction est potentialisée par les kinases tyrosines. Nous avons identifié l'homologue α2-chimérine, comme contenant le même motif de liaison DLP (NPIY143), et confirmé que, comme avec β2- chimérine, ça interagit avec la DLP Dok-4 une fois qu’il est phosphorylé. Aussi, nous avons trouvé que l’interaction avec Dok-4 est grandement amélioré par la conformation active (ouverte) de la β2-chimérine.

3 Fonctionnellement, nous avons trouvé que Dok-4 et β2-chimérine coopèrent pour inhiber l'activité de Rac1 au niveau de la membrane cellulaire et que cet effet inhibiteur nécessite une localisation membranaire médiée par le domaine d'homologie pleckstrin de Dok-4. A partir de ces résultats, nous proposons un modèle par lequel Dok-4 agit pour recruter la β2-chimérine à la membrane, où elle est stabilisée sous sa forme active et est capable d'atténuer la signalisation de Rac1 et, vraisemblablement, des événements de signalisation proliférative en aval.

L’IRI est associée à la progression vers la maladie rénale en phase terminale et l'issue négative du patient. Comprendre le rôle de Dok-4 dans la régulation négative de la signalisation mitogène dans les tubules rénaux blessés aidera à développer des traitements visant à limiter les lésions rénales post-ischémiques et améliorer les résultats des patients. Telles interventions peuvent inclure l'inhibition de l'expression de Dok-4 ou une interférence avec le recrutement ou l'activation de la β2-chimérine.

4 ACKNOWLEDGEMENTS To begin with, I would like to thank my supervisor, Dr. Serge Lemay, for your expertise, your patience and your guidance. I would also like to thank my co-supervisor, Dr. Tomoko Takano, for your continued support throughout this project. I thank you both for all your help.

I would like to thank my thesis committee, Dr. Cristian O’Flaherty, Dr. Suhad Ali, and Dr. Jean-Francois Côté for their interesting and challenging questions, as well as for their helpful suggestions. I would like to thank Dr. R. Lefkowitz, Dr. Morag Park and Dr. N. Lamarche-Vane for generously sharing their plasmids and Dr. Stephane A. Laporte and Yoon Namkung for developing the Rac1 BRET biosensors and helping Cindy Baldwin to set up the assay.

A big thank you to the members of the department of Nephrology, both past and present. You have made my time in the lab enjoyable. A special thanks to Erika Hooker, for welcoming me to the lab and teaching me the ropes. Un remercîment spécial à mes chers compagnons de café, Cindy Baldwin and Lamine Aoudjit. Je vous remercie pour vos conseils techniques, votre encouragement et votre amitié. Il n’y a pas assez de mots pour vous remercier! I would particularly like to thank Cindy for her help with experiments and for her unending patience when it came to answering my interminable questions.

Lastly, I would like to thank my friends and family for their continuous encouragement. To my dearest friend Cindy Audiger, thank you for being a continuous source of inspiration, motivation, support and encouragement. To my sister Alex, thank you for being there to listen to me and for always telling me what I needed to hear, whether or not I wanted to hear it. To my parents, thank you for your support and for always believing in me.

5 Contribution to Original Knowledge This thesis identifies the RacGAPs α2-chimaerin and β2-chimaerin as novel binding partners of the adapter protein Dok-4. We define the biochemical interactions between Dok-4 and α2-chimaerin/β2-chimaerin as requiring the Dok-4 phosphotyrosine binding domain and a phosphorylated tyrosine residue, located at position 143 of α2-chimaerin and position 153 of β2- chimaerin. We have found Dok-4 to cooperate with β2-chimaerin to inhibit Rac1 activity at the membrane. This work identifies Dok-4 as a novel regulator of Rac1 signaling. Of clinical relevance, we found both Dok-4 and β2-chimaerin to be upregulated in the kidney following ischemia-reperfusion injury.

6 Contribution of the Authors In the following monograph, the majority of the experimental data was collected by myself. Cindy Baldwin generated the majority of the constructs used for overexpression and she performed the BRET assays for active Rac and the Rac pull-down assays. Experimental design, analysis and writing were done by myself in collaboration with Dr. Serge Lemay.

7 TABLE OF CONTENTS Abstract……………………………………………………………………………………………2 Résumé…………………………………………………………………………………….………3 Acknowledgements……………………………………………………………………………..…5 Contribution to original knowledge……………………………………………………………….6 Contribution of the authors………………………………………………………………………..7 Table of Contents………………………………………………………………………………….8 List of Figures……………………………………………………………………………………..9 List of Abbreviations…………………………………………………………………………….10

CHAPTER 1: INTRODUCTION, RATIONALE AND OBJECTIVES Introduction………………………………………………………………………………………16 1. EGF Signaling in Ischemic Acute Kidney Injury.…………………………………………...16 1.1. The EGF Receptor………………………………………………...……………………17 1.2. Intracellular Signaling- The Src Family Kinases………………………..………………23 1.3. Rho GTPases and Regulation of Rac1 GTP…………………………………………….26 1.4. Role in IRI ……………………………………………………………………………...28 2. Adapter Proteins in Tyrosine Kinase Signaling...……………………………………………30 2.1. Protein-Protein Interacting Modules: SH2, SH3, PTB Domains.………………...…….32 2.2. Protein-Lipid Interacting Modules: The Pleckstrin Homology (PH) Domain...... ……..33 2.3. Roles of Adapter Proteins in EGFR Signaling ……………………..…………………..34 3. The Downstream of kinase (Dok) family of Adapter Proteins……………..………………..35 3.1. Structure…………………………………………………………………………………36 3.2. Expression……………………………………………………………………………….38 3.3. Function ………………………………………………………………………………...39 3.4. Partners………………………………………………………………………...………..40 3.5. Dok-4……………………………………………………………………………………42 4. Chn Family of Rac-GAPs …………………………………………………………………...43 4.1. Structure…………………………………………………………………………………44 4.2. Expression……………………………………………………………………………….48 4.3. Function ………………………………………………………………………………...49 4.4. Partners………………………………………………………………………………….51 Rationale and Objectives………………………………………………………………………...54

CHAPTER 2: METHODOLOGY AND RESEARCH FINDINGS Methodology …………………………………………………………………………………….56 Research Findings……………………………………………………………………………...... 62

CHAPTER 3: DISCUSSION, FUTURE STUDIES AND CONCLUSIONS Discussion and Future Studies ………………………………………………………………….75 Conclusions……………………………………………………………………………………...81 References……………………………………………………………………………………….82

8 LIST OF FIGURES Figure A: Schematic representation of EGFR/ErbB family receptors and their ligands………..19 Figure B: Schematic Diagram of EGFR Activation…………………………………………….21 Figure C: Potential EGFR signaling pathways activated by ligand binding……………………22 Figure D: Mechanisms involved in activation of Src family kinases…………………………...25 Figure E: The Rho GTPase cycle……………………………………………………………….27 Figure F: Normal repair in ischemic AKI………………………………………………………29 Figure G: Subdivision of adaptor proteins……………………………………………………...31 Figure H: The Dok family proteins…………………………………………………………...... 36 Figure I: The Chimaerin Family Proteins……………………………………………………….44 Figure J: Model for the EGFR-mediated regulation of chimaerins……………………………..47

Figure 1: Interaction of Dok-4 with the Rac1 GAP β2-chimerin-2 (Chn2)…………………….63 Figure 2: Expression of Dok-4 and Chn2 in normal and reperfused kidney…………………....64 Figure 3: The Dok-4 PTB domain interacts with phosphorylated tyrosine 153 of Chn2……….65 Figure 4: Tyrosine 153 of Chn2 is contained within a highly conserved motif in human and murine Chn1 and Chn2…………………………………………………………..66 Figure 5: Dok-4 interacts preferentially with the N-terminal fragment of Chn2 as compared to full-length Chn2……………………………………………………………….67 Figure 6: The Chn2 N-terminal fragment co-localizes with Dok-4……………………………..68 Figure 7: PTB-dependent interaction of Dok-4 with full-length Chn2 is facilitated by the active conformation of Chn2…………………………………………………….70 Figure 8: Measuring the impact of Dok-4 and Chn2 on Rac1 activity………………………….72 Figure 9: Model of Dok-4 anti-proliferative action…………………………………………..…79

9 LIST OF ABBREVIATIONS

A A Alanine aa Amino acid Abl Abelson tyrosine kinase AKI Acute kidney injury Ang II Angiotensin II ARG Amphiregulin Arg Arginine AT1R Angiotensin II type I receptor ATP Adenosine triphosphate

B BCR/Bcr Breakpoint cluster region BiFC Bi-fluorescence complementation Blk B lymphocyte kinase Bmp4 Bone morphogenic protein 4 BRET Bioluminescence resonance energy transfer BTC Betacellulin

C CA Constituently active Caco-2 Human epithelial colorectal adenocarcinoma cells c-Cbl Casitas B-lineage lymphoma CD Cluster of differentiation Cdc42 Cell division control protein 42 Cdk5 Cyclin-dependent kinase 5 cDNA Complementary deoxyribonucleic acid CET Cellules épithéliales tubulaires cGMP Cyclic guanosine monophosphate Chn1 α2-chimaerin Chn2 β2-chimaerin Cip1 Cyclin-dependent kinase inhibitor 1 (alias: p21) CKD Chronic kidney disease CNS Central nervous system Co-IP Co-immunoprecipitation CR Cysteine-rich CRIB Cdc42- and Rac-interactive binding Crk Cyclin-dependent kinase 5 Crm-1 Chromosomal Maintenance 1 Crmp2 Collapsin-response mediator protein-2 Csk C-terminal Src kinase C-terminal Carboxyl-terminus (alias: Carboxy-terminal) CTGF Connective tissue growth factor CXCL12 C-X-C motif chemokine ligand 12 Cys Cysteine

D Dab Disabled

10 DAG Diacylglycerol DGKg Diacylglycerol kinase g DLP Domaine de liaison à la phosphotyrosine DNA Deoxyribonucleic acid Dok Downstream of kinase

E E Glutamic acid EGF Epidermal growth factor EGFR Epidermal growth factor receptor EGFP Enhanced green fluorescent protein EGN Epigen Elk ETS domain-containing protein EphA erythropoietin-producing human hepatocellular receptor A EPR Epiregulin ErbB Erythroblastic oncogene B ERK Extracellular signal-regulated kinase ESRD End-stage renal disease

F F Phenylalanine FC Facteurs de croissance FGF Fibroblast growth factor Fgr Gardner-Rasheed feline sarcoma viral (v-fgr) oncogene homolog FRS2 Fibroblast growth factor receptor substrate 2 beta Fyn Proto-oncogene tyrosine-protein kinase Fyn

G G Glycine G0 G zero phase G1 Gap 1 phase Gab Grb2-associated binder GAP GTPase-activating protein GDI Guanine nucleotide dissociation inhibitor GDNF Glial cell line-derived neurotrophic factor GDP Guanine diphosphate GEF Guanine nucleotide exchange factor GFP Green fluorescent protein GPCR G-protein-coupled receptors Grb2 Growth factor receptor-bound protein 2 GST Glutathione S-transferase GST-PD Glutathione S-transferase pull down assay GTP Guanine triphosphate

H h Hour H Histidine HB-EGF Heparin-binding EGF-like growth factor hChn1 human α2-chimaerin hChn2 human β2-chimaerin

11 Hck Hematopoietic cell kinase HEK Human embryonic kidney HeLa Cell line derived from cervical cancer cells taken from Henrietta Lacks HER Human epidermal growth factor receptor HGF Hepatocyte growth factor His Histidine

I I Isoleucine IgM Immunoglobulin M IGF Insulin-like growth factor IL-2 Interleukin 2 IP Immunoprecipitation IR Insulin receptor IRI Ischemia-reperfusion injury/ insulte d'ischémie-reperfusion IRS Insulin receptor substrate

J JAK Janus kinase JNK c-Jun N-terminal kinase (alias: SAPK)

K kDa Kilodalton

L L Leucine Lck Lymphocyte-specific protein tyrosine kinase LR1G1 Leucine rich repeats and immunoglobulin-like protein 1 Luc Luciferase Lyn Lck/Yes-related novel protein tyrosine kinase

M M Methionine MAPK Mitogen-activated protein kinase MCF-7 Michigan Cancer Foundation-7 cell line mChn1 mouse α2-chimaerin mChn2 mouse β2-chimaerin M-CSF Macrophage colony-stimulating factor MDCK Maden-Darby canine kidney MEK MAPK/ERK kinase Mig-6 Mitogen-inducible gene 6 (alias: RALT, Gene 33) miRNA microRNA min Minute mL Millilitre MLK2/3 Myosin light chain kinase 2/3 mM Millimolar mRNA Messenger RNA MuSK Muscle-specific tyrosine-protein kinase receptor Myr Myristoylated

N N Asparagine

12 NES Nuclear export signal ng Nanogram NGF Nerve growth factor N-lobe Amino-terminal lobe NMDA N-methyl-D-aspartate NRG Neuregulin NR2A NMDA receptor subunit 2A NT Amino-terminal (alias: N-terminal) N-terminal Amino-terminal

P P Proline p21 Cyclin-dependent kinase inhibitor 1 (alias: Cip1, Waf1) p23 Transmembrane Protein Tmp21 p35 Cyclin-Dependent Kinase 5, Regulatory Subunit 1 p38 Mitogen-activated protein kinases PA Phosphatidic acid PAK p21-activated kinase PC12 Cell line derived from a pheochromocytoma of the rat adrenal medulla pcDNA Plasmid cytomegalovirus promoter deoxyribonucleic acid vector PDGF Platelet-derived growth factor PE Phorbol ester PH Pleckstrin homology PI3K Phosphoinositide-3-kinase (alias: phosphatidylinositol-3 kinase) PI(3,4)P2 Phosphatidylinositol 3,4-diphosphate PI(4,5)P2 Phosphatidylinositol 4,5-bisphosphate PI(3,4,5)P3 Phosphatidylinositol 3,4,5-trisphosphate PKB Protein Kinase B PKC Protein Kinase C PKD Protein Kinase D PLC Phospholipase C PM Plasma membrane PMA Phorbol myristate acetate pRB Retinoblastoma family protein P-Rex1 Phosphatidylinositol-3,4,5-trisphosphate-dependent Rac exchange factor 1 PS Phosphatylserine PTB Phosphotyrosine binding PTK Protein tyrosine kinase PTPN1 Protein tyrosine phosphatase 1B (alias: Tyrosine-protein phosphatase non- receptor type 1) pTyr Phosphotyrosine pY Phosphotyrosine

R R Argenine RALT Receptor-associated late transducer (alias: Mig-6, Gene 33) RNA Ribonucleic acid

13 RSV Rous sarcoma virus RTK Receptor tyrosine kinase

S S Synthesis phase SFK Src family kinases SH1 Src homology 1 SH2 Src homology 2 SH3 Src homology 3 Shh Sonic hedgehog SHIP-1 SH2 domain-containing inositol phosphatase 1 siRNA Short interfering ribonucleic acid (alias: Silencing RNA) SOCS Suppressor of cytokine signaling Sos Son of sevenless SRE Serum response element Srf Serum response factor STAT Signal transducers and activators of transcription Syk Spleen tyrosine kinase

T TGF-a Transforming growth factor a TKD Tyrosine kinase domain TM Transmembrane Trk Ttropomyosin-related kinase TRMP2 Transient receptor potential cation channel subfamily M member 2 (alias: Transient receptor potential melastatin 2) TSS Transcription start sites Tyr Tyrosine

U uL Microlitre ug Microgram

V V Valine VEGF Vascular endothelial growth factor Ver Version

W Waf1 Cyclin-dependent kinase inhibitor1 (alias: Cip1, p21) WT Wild-type

X X Any amino acid

Y Y Tyrosine Y2H Yeast two-hybrid Yrk Yes-related kinase

CHAPTER 1: INTRODUCTION AND OBJECTIVES

15 Introduction

The epidermal growth factor receptor (EGFR), a versatile signal transducer, is an important cell surface receptor with intrinsic protein tyrosine kinase (RTK) activity that has been well conserved during evolution [1]. Tyrosine phosphorylation, such as that resulting from EGFR activation, is a highly regulated post-translational modification, and a fundamental mechanism of intracellular signal transduction and regulation in all eukaryotic cells [1, 2]. Tyrosine kinases are essential in nearly all aspects of cell life, regulating signaling pathways and cellular processes that mediate metabolism, transcription, cell-cycle progression, differentiation, development, cytoskeleton arrangement and cell movement, apoptosis, intercellular communication, and neuronal and immunological functions [2].

Aberrant tyrosine phosphorylation underlies many human diseases, including cancer [4], and tyrosine kinase signaling plays an important role in mediating the cellular processes underlying ischemic acute kidney injury [5]. Currently, 47 protein kinase inhibitors, 42 of which target tyrosine kinases, are approved for clinical use in the treatment of various cancers [6], highlighting the importance of understanding the structure and function of these signaling molecules.

1. EGF Signaling in Ischemic Acute Kidney Injury Acute kidney injury (AKI) has been recognized as a major public health burden [7] and an important independent risk factor for developing chronic kidney disease (CKD) and hastening the development of end-stage renal disease (ESRD) [5], morbidity and mortality. The global population prevalence of CKD is over 10%, rivaling that of diabetes, and increases with age to over 20% in patients over 60 years and over 35% in those older than 70 years [7].

Renal ischemia-reperfusion injury (IRI) is a major cause of AKI in both native and transplanted kidneys and comprises two phases. Initially, during ischemia, a generalized or local impairment in blood flow to the kidneys results in a deficiency in oxygen, ATP and nutrient delivery to and waste product removal from cells of the kidney [8]. Subsequently, during

16 reperfusion, the reestablishment of blood flow to the kidney is linked to the activation of inflammatory pathways, associated with the generation of free radicals and reactive oxygen species, the production of cytokines from and the infiltration of immune cells into the kidney tubules [5], [9]. Renal ischemia-reperfusion causes tissue damage, particularly to the epithelial cells of the proximal tubules of the outer medulla and leads to functional impairment of the kidney.

Since AKI often occurs unexpectedly and rapidly, the window for prevention and early therapy is very narrow. Therefore, there is an obvious need for developing therapeutic interventions for enhancing reparative processes in established AKI. Multiple studies have demonstrated a role for exogenous growth factor-induced EGFR activation in facilitating and accelerating tubular repair in rodent models of AKI [10]–[14]. Disappointingly, these results failed to translate in porcine models [15] and human clinical trials of AKI [16], [17]; however, it may be premature to abandon investigating the impact of EGFR activation on AKI outcome in humans, as the failure of previous clinical trials may have been impacted by flawed experimental design [18], which can be avoided in future trials. Furthermore, since sustained activation of EGFR has been shown to lead to renal fibrogenesis in a murine model of AKI [19], it is possible that endogenous regulatory mechanisms may be attenuating the efficacy of growth factor-based interventions by negatively regulating EGFR signaling; therefore, a better understanding of these negative regulatory mechanisms would provide key insight into maximizing the therapeutic value of EGFR signaling in AKI.

1.1. The EGF Receptor

The EGFR (also known as HER-1; ErbB-1) is a 170 kDa transmembrane protein, belonging to the family of receptor tyrosine kinases (RTKs), cell-surface receptors with intrinsic protein tyrosine kinase (PTK) activity [1]. Playing an important role in initiating epithelial cell signaling cascades, the EGFR is a prime example of receptor-mediated activation of cellular proliferation, survival, and differentiation in both development and normal physiology, as well as in pathophysiological conditions [20]. EGFR is one of four transmembrane growth factor receptor

17 proteins that share similarities in structure and function. Other members of the ErbB/HER family include HER2/neu (ErbB-2), HER3 (ErbB-3), and HER4 (ErbB-4) [21].

The overall architecture of EGFR is shared by many other receptors, including those for insulin, platelet-derived growth factor (PDGF), fibroblast growth factor (FGF), and vascular endothelial growth factor (VEGF), which are collectively known as RTKs [22]. Structurally, the EGFR comprises an extracellular ligand-binding domain, a single membrane- spanning region, a cytoplasmic protein tyrosine kinase domain, and a C-terminal tail with multiple phosphorylation sites [20].

The extracellular ligand-binding region of EGFR is comprised of four subdomains arranged as a tandem repeat of two types of domains. The first and third domains are homologous to one another and are designated domains I and III or L1 and L2, respectively; the second and fourth domains are also homologous to one another and are designated domains II and IV or CR1 and CR2, respectively [23]–[25], where CR represents cysteine-rich regions, where nearly 50 conserved cysteines are found in these two domains. Studies using mutant and chimeric EGF receptors and receptor fragments demonstrated that ligand binding is mediated primarily by domain III with some contribution from regions on domain I [24], [26]. The transmembrane domain of EGFR is comprised of single alpha-helix and the cytoplasmic domain contains a juxtamembrane cytoplasmic domain, a Src homology (SH1) tyrosine kinase domain (TKD), and a carboxy-terminus tail with multiple autophosphorylation sites that binds to proteins containing phosphotyrosine binding Src homology 2 (SH2) or phosphotyrosine binding (PTB) domains [20], [27] (Figure A).

18 A 1 B EGF Domain I TGF-α HB-EGF 165 ARG EPR NRG1 NRG3 EGN BTC NRG2 NRG4 Domain II 310 Domain III

480 EGFR ErB2 ErB3 ErB4 Domain IV 620 TM PM 643 685

TKD 953 I Y I Y I Y I Y Y I 1186

Y Y Y Y YY Y Y YY Y Y YY Y Y YY Y Y

Figure A: Schematic representation of EGFR/ErbB family receptors and their ligands. (A) The domain composition of human EGFR is shown. The extracellular region contains four domains: Domain I (amino acids 1– 165), domain II (amino acids 165–310), domain III (amino acids 310–480), and domain IV (amino acids 480– 620). Domains I and III are closely related in sequence, as are domains II and IV. Shown are representations of the structures of domains I and IV. The ~23-amino-acid transmembrane (TM) domain spans the plasma membrane (PM). A representative EGFR tyrosine kinase domain (TKD) structure is shown, followed by a largely unstructured c-terminal tail (amino acids 953–1186) that contains at least five tyrosine autophosphorylation sites. (B) EGFR is one of four members of the EGFR/ErbB family in humans. The other members are ErbB2/HER2, for which no soluble activating ligand is shown; ErbB3/HER3, which has a significantly impaired kinase domain; and ErbB4/HER4. EGFR is activated by the EGFR agonists: EGF itself, TGF-α (transforming growth factor α), ARG (amphiregulin), and EGN (epigen). The bispecific ligands regulate both EGFR and ErbB4: HB-

19 EGF (heparin-binding EGF-like growth factor), EPR (epiregulin), and BTC (betacellulin). Neuregulins (NRGs) 1 and 2 regulate ErbB3 and ErbB4, whereas NRG3 and NRG4 appear to be specific for ErbB4. Adapted from: Lemmon, M.A., 2014 [27].

EGFR is regulated by at least eight different soluble activating ligands in humans: the EGF protein itself, transforming growth factor a (TGF-a), betacellulin (BTC), heparin-binding EGF-like growth factor (HB-EGF), amphiregulin (ARG), connective tissue growth factor (CTGF), epiregulin (EPR), and epigen (EGN) [20], [28]. Each of the mature peptide growth factors is characterized by an EGF-like domain that is responsible for receptor binding and activation, with a characteristic pattern of six spatially conserved cysteines that form three intramolecular disulfides. The EGFR ligands are all produced as membrane-bound precursor proteins [28], which are cleaved by cell-surface metalloproteases to yield the active growth factors [29].

EGFR activation mediates many signaling processes through EGFR “transactivation.” The unbound EGFR is intrinsically autoinhibited and transactivation occurs either through activation of metalloproteinase-dependent ligand release or through non-ligand-dependent receptor activation [20]. Upon ligand binding to a single EGFR molecule, the bound EGFR will either form an asymmetric homodimer with an additional EGFR molecule or a heterodimer with another member of the ErbB family. EGFR has also been reported to heterodimerize with receptors for hepatocyte growth factor (HGF), insulin-like growth factor (IGF)-I, and PDGF [30], the biologic significance of these interactions remains to be elucidated. The EGFR dimer is allosterically activated when the C-lobe of one TKD is juxtaposed against the N-lobe of the other TKD, leading to activation of tyrosine kinase activity [31], [32]. The active tyrosine kinase domains of both members of the receptor dimer auto-phosphorylate specific tyrosine residues at positions 1068, 1148, 1173, 992 and 1086 [33]–[36] in the cytoplasmic tail, providing high- affinity binding sites for subcellular signaling protein complexes [32] (Figure B). Multiple signaling pathways are activated by EGFR depending on the activating ligand, including the mitogen-activated protein kinase (MAPK) cascades of Ras/Raf/MEK/ERK, p38, JNK, Jak-STAT and Shc/Grb2/Sos1/Rsk2; the survival associated phosphoinositide 3-kinases (PI3K)/Akt

20 pathway; the proto-oncogenic Src; the small GTPases such as Rho and Rac; and the serine/threonine kinase pathway phospholipase C (PLC)-g /Ca2+/ protein kinase C (PKC) and protein kinase D (PKD) [20] (Figure C).

Inactive (monomer or dimer) Active dimer

EGF Extracellular domain

N-lobe Activator Activation loop TKD P Activator P C-lobe P P C-terminal tail Receiver Receiver P P P Asymmetric dimer P P P

P Autophosphorylation sites

Figure B: Schematic Diagram of EGFR Activation. Activation of EGFR by EGF results in the formation of an asymmetric kinase domain dimer; Plasma membrane (PM), Tyrosine kinase domain (TKD), phosphorylation site (P). Adapted from: Jura, N., 2009. [31]

21 EGFR

PM DAG P Shc P PLCγ PKC Rsk2 Sos1 Grb2 IP3 P Ca2+ Ras P Src Vav Raf P P Rac-GTP FAK

MEK Nck Grb2 Gab1 Jak1/2 STAT1/3 ERK1/2 PAK PI3K

JNKK

JNK Akt

Figure C: Potential EGFR signaling pathways activated by ligand binding. Following binding, a number of cytoplasmic tyrosine residues are autophosphorylated by the intrinsic receptor kinase. Major pathways mediating EGFR actions include Shc/ Grb2/Sos1/Rsk2 and Ras/Raf/ MEK/ERK, the JNK pathway, PLC/DAG/PKC, the Src pathways, the Jak/STAT pathways and PI3K/Akt in addition to activating small GTPases, such as Rac. Adapted from: Chen, J., 2016 [20]

As a consequence of mediating multiple cellular events, EGFR signaling is tightly regulated at both the level of the receptor and its ligands, this regulation is differential, depending on the specific activating ligand. For example, like all RTKs, the regulation of EGF-, HB-EGF- and BTC-bound EGFRs involves a two-step process of endocytosis [37], [38], initially involving a ligand-dependent internalization step, whereby activated EGFRs cluster in both clatherin-coated and non-clatherin-coated pits where they are immediately internalised into early endosomes [39], [40], removing them from the cell membrane; followed by targeting of the internalized receptors to lysosomes where they undergo ubiquitinylation and degradation, the

22 latter involving c-Cbl ubiquitin E3 ligase, recruited through EGFR auto-phosphorylation [41]. In contrast, TGF-a- and EPR-binding EGFRs induces complete recycling from endosomes, which has been suggested to be due to increased dissociation of the ligands from the receptor in acidic endosomes[42], and deactivation of the EGFR through dephosphorylation by the recruitment of protein tyrosine phosphatases such as PTPN1/SHP2 [43], leading to recycling of the deactivated EGFR to the plasma membrane.

Multiple other proteins have also been identified to act as feedback inhibitors of EGFR signaling. Mitogen-inducible gene 6 (Mig-6)/receptor-associated late transducer (RALT)/Gene 33 is an early-response gene induced by EGF, whose transcription is regulated by ERK signaling. Mig-6 binds the EGFR TKD, inhibiting its kinase activity, and EGF signaling [44]. Fibroblast growth factor receptor substrate 2 beta (FRS2), constitutively binds to EGFR, regardless of ligand stimulation. After activation of ERK by various growth factors, including EGF, the phosphorylated ERK binds to FRS2 and inhibits EGFR autophosphorylation and signal transduction [45]. Suppressor of cytokine signaling 3 (SOCS3)/SOCS4/SOCS5 is an early- response gene induced by EGF. SOCS proteins bind to autophosphorylated EGFR via their Src homology 2 (SH2) domains and recruit E3 ubiquitin ligase complex via their SOCS domains, leading to the ubiquitinylation and degradation of EGFR [46], [47]. Immunoglobulin-like domains 1 (LRIG1) is another early-response gene induced by EGF. It binds to EGFR via its extracellular domain and promotes Cbl-mediated ubiquitination, followed by internalization and degradation of EGFR [48]. LRIG1 is also cleaved by ADAM17, and the soluble extracellular domain can inhibit ligand binding to EGFR [49], without physical downregulation of the receptor. Studies in cancers suggest a role for microRNAs (miRNAs) in regulating EGFR activity, either through direct modulation of the receptor itself [50], [51] or by acting on the EGFR-associated proteins which regulate the EGFR [52]. These negative regulatory mechanisms are essential for proper EGFR signaling as dysregulation of the signaling pathways can result in uncontrolled cellular proliferation leading to malignant transformation and cancer progression.

1.2. Intracellular Signaling- The Src family kinases (SFKs)

23 The origins of the SFKs are over a century old, beginning with the pioneering studies of Peyton Rouse who found that a fibrosarcoma could be transmitted between chickens in a cell- free extract of the tumor [53]. The transmissible agent, Rous sarcoma virus (RSV), was found to transform cells by virtue of the presence within its genome of a viral oncogene, v-src, which is derived from a normal cellular gene that has been picked up, or transduced, by the virus [54]– [56]. The SFKs make up the largest family of non-receptor tyrosine kinases, playing important roles in regulating signals from various cell surface receptors, including EGFR, in different cellular contexts. Comprised of nine members, divided into two subgroups: the ubiquitously expressed Src-related subgroup, comprising c-Src, Yes, Fyn and Fgr while the Lyn-related subgroup is made up of Lyn, Hck, Lck, Blk and Yrk [57], [58].

Structurally, all SFKs share a conserved domain arrangement consisting of a variable N- terminal region of 50-70 residues [59], unique to each SFK member but always containing a conserved myristoylation or palmitoylation site [60], [61]; followed by the ∼50 amino acid Src homology 3 (SH3) domain which directs specific association with proline rich motifs related to the core PXXP consensus [62]. Next is the ∼100 amino acid Src homology 2 (SH2) domain which provides interaction with phosphotyrosine motifs, with SKFs showing highest affinity for the consensus sequence pYEEI [62]. The last domain is the C-terminal bilobal Src kinase domain (∼300 residues), or Src homology 1 (SH1), responsible for the enzymatic activity [57] and a negative regulatory tail [63].

Under basal conditions, SFKs are autoinhibited via two intramolecular interactions, whereby phosphorylation of Tyr527 (Src), in the C-terminal tail, by the cytoplasmic tyrosine kinase C- terminal Src kinase (Csk), binds the SH2 domain [64]; and the SH3 domain interacts with sequences in the catalytic domain, as well as with proline-rich sequences in the linker region that lies between the SH2 and catalytic domains [65], [66]. An auto-phosphorylation site, corresponding to Tyr416 (Src), contained within a classical kinase activation loop, is phosphorylated to induce a conformational change allowing the kinase to assume an active conformation [63]. The inhibitory intramolecular interactions of the SH2 and SH3 domains are released by phosphorylation of Tyr416 [57], by dephosphorylation of Tyr 527 by a tyrosine

24 phosphatase [63], or by competition for the SH2 and SH3 domains by high-affinity ligands such as active RTKs [58], ultimately leading to SFK activation (Figure D).

Inactive Kinase Activated Kinase Y416 phosphorylation

pY527 dephosphorylation

SH3 SH3 N-lobe SH2 P Y416 Activation Kinase SH2 pY527 loop C-terminal tail C-lobe

Kinase P pY416 P Phosphorylated tyrosine pY527

C-terminal tail

Figure D: General structural schematic of the Src family kinases in their inactive and active configurations. The left panel shows a model of the structure of inactivated Src family kinases, where the SH2 domain interacts with the phosphorylated C-terminal tyrosine (pY527 in this model of Src), the SH3 domain interacts with the SH2-kinase connector and the dephosphorylated activation loop (Y416) folded back over the substrate binding site. The right panel represents a model for the activated state of Src in which the intramolecular interactions of the SH3 and SH2 domains are disrupted, the C-terminal tyrosine is dephosphorylated (Y527), and the activation loop is phosphorylated (pY416) and is folded away from the substrate binding site and allows the two kinase lobs (N and C) to form a kinase competent catalytic cleft. PM: plasma membrane. Adapted from: Thomas, S.M., et al., 1997 [63].

25 There is considerable evidence of bidirectional interplay between SFKs and the EGFR, both are overexpressed in many types of tumors [67] and c-Src, in particular, has been shown to potentiate the mitogenic and oncogenic effects mediated by EGFR [68]. Downstream of EGFR activation and auto-phosphorylation, Src kinases bind to phosphorylated tyrosines on the intercellular domain of the activated receptor through their SH2 domain, leading to their full activation and propagation of downstream signalling cascades, acting on a variety of substrates, regulating gene transcription, cell adhesion and spreading, focal adhesion dynamics, formation of lamellipodia, actin cytoskeleton rearrangements and motility, cell survival and differentiation [63], [64], [69]. Alternatively, SFKs, such as c-Src, can activate the EGFR by phosphorylating its cytoplasmic Tyr845, potentiating receptor signaling even in the absence of ligand stimulation [70].

Three SFKs, Src, Fyn and Lyn are expressed in renal epithelial cells [12]. During renal regeneration following AKI, SFKs are implicated in mediating renal proximal tubule cell dedifferentiation, through activation of the EGFR/PI3K signaling pathway, in addition to promoting proximal tubule cell proliferation and migration [12].

1.3. Rho GTPases and Regulation of Rac1 GTP

Rho GTPases, belonging to the Ras superfamily of small GTPases, are molecular switches that control a wide variety of signal transduction pathways that regulate diverse cellular processes in eukaryotic cells. The most extensively studied Rho GTPases are RhoA, Rac1, Rac2, and Cdc42. Rac1, like all members of the Ras superfamily, functions as a conformational switch by cycling between active GTP- and inactive GDP-bound forms. The cycle is regulated by two classes of protein: guanine nucleotide exchange factors (GEFs), which catalyze nucleotide exchange, resulting in the activation of Rac1 due to the higher concentration of GTP than GDP in cells; and GTPase-activating proteins (GAPs), which stimulate GTP hydrolysis, leading to the inactivation of Rac1. Additionally, Rac1 is regulated by guanine nucleotide dissociation inhibitors (GDIs), which extract and sequester GDP-bound forms from the membrane to the cytosol and inhibit the release of GDP from Rac1 [71], [72] (Figure E). GTP- Rac1 is able to bind a variety of downstream effectors, which initiates a variety of cell-type specific responses,

26 such as actin cytoskeleton rearrangement, microtubule dynamics, cell adhesion and polarity [71], cell migration, gene transcription and cell proliferation [73], [74].

GEF Rac1 Rac1 GDP GTP inactive active

GAP

P GDI Rac1

inactive

Effectors

Figure E: The Regulation of Rac1. Rac1 cycles between an active (GTP-bound) and an inactive (GDP-bound) conformation. In the active state, it interacts with one of its target proteins (effectors). The cycle is highly regulated by three classes of protein: in mammalian cells, guanine nucleotide exchange factors (GEFs) catalyze nucleotide exchange and mediate activation; GTPase-activating proteins (GAPs) stimulate GTP hydrolysis, leading to inactivation; and guanine nucleotide exchange inhibitors (GDIs) sequester the inactive GTPase in the cytosol. Adapted from: Etienne-Manneville, S. and Hall, A., 2002 [71].

Rac1 is a ubiquitously expressed 21 kDa protein, that is activated by a variety of stimuli, including growth factors such as EGF and G-protein-coupled receptor ligands. Tyrosine-kinase receptors may convey signals to Rac-GEFs via intermediate molecules, such as the class I PI3K, as described for P-Rex1 [75], whereas PI3K-independent regulation of GEFs may involve their direct binding to and phosphorylation by the tyrosine-kinase receptor, as described for Vav2 with EGFR [76]. Active Rac1 interacts with specific effectors through domains that coordinate activation of a multitude of signaling cascades that influence diverse physiological outcomes,

27 particularly involving roles in the control of actin cytoskeleton organization, migration, metastasis, transformation, gene expression and cell-cycle progression [77]. The family of p21 activating kinases (PAK) was among the first described Rac1 effector proteins. PAK binds Rac1 in a GTP-dependent manner, stimulating PAK kinase activity and leading to altered cytoskeletal dynamics, adhesion, and transcription [78], [79]. Rac1 signals through PAK to activate c-Jun N- terminal kinase (JNK) and p38 families [80], to regulate gene expression. In addition, Rac1, through PAK, can influence transmembrane guanylyl cyclase activity and the second messenger cGMP production [81] and mediates canonical JNK regulated Wnt-signaling to the T-cell factor transcription factor [82]. Rac1 has also been shown to influence nuclear signaling through its effectors MLK2/3, which have been shown to activate the JNK pathway [83], [84]. Rac1 signaling can be important for cellular transformation via modulation of anti-apoptotic and cell cycle machineries. Rac1 positively regulates transcription at NFkB transcription factor- dependent promoters [85] and facilitates PI3K-dependent activation of Akt serine/threonine kinase [86], [87], thereby permitting the survival of transformed cells. Rac1 can also influence proliferation and transformation through regulation of cyclin D1, a cell cycle protein that is frequently overexpressed in cancer [80], [88], and the cell-cycle inhibitor p21Cip1/Waf1 [73]. Thus, Rac1 is a critical component of a complex EGFR signaling network, given its ability to directly or indirectly impact a vast array of cellular signals.

Overall, Rac plays essential roles in the control of actin cytoskeleton organization, migration, metastasis, transformation, gene expression and cell-cycle progression [3–6].

Since renal IRI is associated with prominent alterations in cell proliferation and cytoskeletal organization, it is expected that Rac and other Rho GTPases play important roles during its course. Indeed, in a mouse model of renal IRI, the nonselective transient receptor potential cation channel subfamily M member 2 (TRPM2) promoted Rac1 activation, with active Rac1 physically interacting with TRPM2 at the cell membrane; inhibition of Rac1 reduced oxidant stress and ischemic injury in vivo, demonstrating that TRPM2-dependent Rac1 activation increases oxidant stress and identifying TRPM2 and Rac1 as potential therapeutic targets in reducing ischemic kidney injury [89].

1.4. Role in IRI

28 EGFR has been most predominantly studied in the context of oncogenesis, however its role in renal ischemic AKI is becoming clear. AKI, is defined as an abrupt and sustained reduction in kidney function, measured by a rise in serum creatinine and blood urea nitrogen or a fall in urine output. AKI can arise in a variety of clinical situations, such as ischemia/reperfusion, sepsis, trauma and nephrotoxin exposure [13], highlighting the need for a better understanding of the causes and complications of AKI [5].

Following ischemic injury, the polarized epithelial cells of the proximal tubules are subject to disruption of the actin cytoskeleton and a loss of cell polarity [5]; there is a loss of cell-cell contacts and an increase in intercellular permeability [90]. Some tubule epithelial cells undergo cell death by apoptosis and necrosis [5] and are sloughed off into the tubule lumen. Yet, in contrast to the heart or brain, the kidney can completely recover from an ischemic insult that results in cell death. During recovery from IRI, surviving tubular epithelial cells migrate, dedifferentiate and rapidly proliferate, replacing the irreversibly injured tubular epithelial cells and restoring tubular integrity and kidney function [5], [91] (Figure F).

Normal polarized Ischemia-reperfusion tubular epithelium

Loss of polarity/ Loss of brush border Differentiation/ Necrosis Apoptosis Reestablishment of polarity

Cell death Proliferation

Migration and dedifferentiation of Sloughing off of dead and viable cells viable cells Adhesion molecules Figure F: Normal repair in ischemic acute kidney injury (AKI). The current understanding of tubular injury and repair after ischemic AKI. Following IRI, the normally highly polar epithelial

29 cell loses its polarity and brush border and there is aberrant localization of proteins on the cell membrane. Injured epithelial cells undergo cell death by either necrosis or apoptosis. Some of the injured viable cells and cell necrotic cell debris is released into the lumen, exposing the basement membrane. Viable epithelial cells dedifferentiate and migrate to cover denuded areas of the basement membrane. These cells undergo division and replace lost cells. Ultimately, the cells differentiate and reestablish the normal polarity of the epithelium. Adapted from: Bonventure, J.V. and Yang, L., 2011 [5].

EGFR signaling plays important roles in renal repair after acute injury. Activation of EGFR, through administration of exogenous EGF, enhances renal tubule cell regeneration and repair and accelerates the recovery of renal function in IRI-induced AKI [92]. Renal pro-EGF mRNA and EGF excretion decrease post-IRI [93]; however, other EGFR ligands, particularly HB-EGF, are upregulated in AKI models and appear to mediate renal epithelial cell proliferation [94], [95]. In addition, both EGFR expression [96] and activation increase in response to AKI [97], [98], and is required for renal proximal tubule epithelial cell proliferation, mediated by PI3K [14], migration, mediated by p38 kinase [14], dedifferentiation, mediated by both PI3K and p38 [14], [99] and redifferentiation [97]. In waved-2 mice, a mouse strain with a point mutation in the EGFR that results in a 90% reduction in receptor tyrosine kinase activity, renal dysfunction and histological damage was more severe and functional recovery was much slower after folic-acid-induced AKI [11]. Similarly, renal functional recovery was significantly delayed following ischemia and reperfusion injury in mice with a specific EGFR deletion in the renal proximal tubule or in mice treated with erlotinib, a specific EGFR tyrosine kinase inhibitor [100]. These data provide strong evidence that EGFR is critically involved in promoting kidney recovery from acute injury.

2. Adaptor Proteins in Tyrosine Kinase Signaling It is well known that a group of proteins, called adaptor proteins, play an essential role in the propagation of tyrosine kinase signaling. Adaptor proteins are multi-modular proteins that mediate both protein-protein interactions and protein-lipid interactions, they may also contain motifs encompassing tyrosine phosphorylation sites and are defined by their lack of enzymatic or transcriptional activity. Adaptor proteins play a central role in the integration of intracellular

30 signal transduction, acting as docking proteins, anchoring signaling complexes to particular subcellular structures, such as the cell membrane, or acting as scaffolding proteins, which bring together many different signaling proteins in complex [101] (Figure G).

Adaptor

Docking Scaffolding

A A A A

B B B B

C C C C

A Protein interacting module of adaptor protein

A,B,C Any molecules

Membrane-bound docking site

Signaling-dependent interaction Constitutive interaction

Figure G: Subdivision of adaptor proteins. Adaptor proteins are molecules of multimodular structure without enzymatic activity that link several proteins together or recruit proteins or protein complexes to subcellular structures, utilizing both constitutive and temporary interactions. Docking and scaffolding proteins can be viewed as subsets of the adaptor superfamily. Docking proteins are specialized to dock one or more interacting partners to subcellular structures. Scaffolding proteins can bind multiple signaling proteins, bringing together members of enzymatic cascades, and often organizing large signaling complexes localized to specific subcellular structures. Adapted from: Csiszar, A. 2006 [102].

Adaptor proteins are important in conferring signaling specificity and controlling signal duration. By engaging different combinations of adaptors, one signal can produce different outcomes. Distinctive phosphorylation sites, mediating the interaction or dissociation of particular effector molecules, confers an additional level of signal specificity. Additional post-

31 translational modifications, such as sumoylation, methylation and acetylation can contribute to adaptor protein regulation [102]. To carry out their functions, adaptor proteins are composed of different small modules of 50-200 amino acids [103], that direct their constitutive and/or signal- regulated association with other proteins, with membranes, and with other cellular components. SH2, Src homology 3 (SH3) and PTB domains, recognize phosphorylated tyrosine (pTyr) residues and are important for mediating protein-protein interactions, while pleckstrin homology (PH) domains mediate adaptor protein binding to lipids [102].

2.1. Protein-Protein Interacting Modules: SH2, SH3, PTB Domains

The SH2 domain was the first signaling module identified to possess specificity for pTyr residues [104]. In addition to adaptor proteins, SH2 domains are present in a diverse array of molecules, including kinases, phosphatases, GTPases and transcription factors [103], [105], emphasizing their importance in tyrosine kinase signaling. Structurally, the SH2 domain is a compact 100 amino acid residue molecule, containing a conserved FLVR motif [106], [107]. The three-dimensional structure of SH2 domains is highly conserved, composed of two a-helices and an anti-parallel b-sheet, made up of three to four b-strands. These structures form a positively charged binding pocket, utilizing a critical argenine residue, required for binding pTyr of target ligands. In canonical binding, the residues surrounding the critical Arg typically engage amino acids from position +1 to +6 (C-terminal of the pTyr) of the ligand, and it is this sequence that dictates the specificity of a given SH2 domain [102], [108].

In contrast to SH2 domains, PTB domains are the only modules exclusively present in adaptor proteins. They are slightly larger than SH2 domains, composed of about 200 amino acids. PTB domains have a highly conserved 3-D β-sandwich structure, termed the pleckstrin homology (PH) domain “superfold” [109], comprised of seven anti-parallel b-strands forming a sandwich of two orthogonal b-sheets capped by a C-terminal a-helix of variable length. Generally, peptide binding is mediated by residues in the PTB b5 strand and the C-terminal a- helix [110], [111]. Typically, peptide binding by the PTB domain is pTyr-dependent, involving an NPXpY motif (where X is any amino acid), considered the canonical binding motif for PTB- domain-containing proteins; however, many PTB domains can bind their target proteins independent of ligand tyrosine phosphorylation. Additionally, PTB domains commonly bind

32 phospholipid acidic head groups, which helps to localize them to membrane or juxtamembrane regions where they can easily bind RTKs and mediate downstream signaling. Binding to phospholipid head groups is mediated by a positively charged binding pocket composed of a cluster of basic residues and is a distinct event from the PTB domain-containing protein’s interactions with pTyr motifs [111].

Using structural, functional and evolutionary data, PTB- domain-containing proteins are classified into three groups: pTyr-dependent Shc-like; pTyr-dependent insulin receptor substrate (IRS)-like and pTyr-independent Dab-like [111], which are distinct in their ability to bind a phosphorylated NPXY motif. The Shc- and IRS-like modules preferentially bind phosphorylated substrates, whereas Dab-like PTB-domain-containing proteins preferentially bind to residues that are unphosphorylated or in which the tyrosine is replaced with phenylalanine [112].

In addition to phosphotyrosine-binding domains, modules functioning independently of tyrosine kinase activity exist that are frequently part of tyrosine kinase signalling-linked adaptors. SH3 domains, for example, are small protein modules of about 60 amino acids that bind to proline-rich recognition sequences [113].

Protein-protein interactions are fundamental in many signaling cascades, making SH2, SH3 and PTB domains essential components of a large majority of signal transduction pathways. The binding interactions mediated by these domains allow for the formation of numerous types of specific, transient and often signal-dependent protein complexes, important in the cellular responses to a variety of stimuli.

2.2. Protein-Lipid Interacting Modules: The pleckstrin homology (PH) Domain

PH domains are protein-lipid interaction modules, approximately 100-120 amino acid residues long, involved in signal propagation and subcellular localization. They have been identified in nearly 100 different eukaryotic proteins, many of which participate in cell signaling and cytoskeletal regulation, including serine/threonine and tyrosine kinases, regulators of small

33 G-proteins, endocytic GTPases, adaptor proteins, lipid-associated enzymes, and cytoskeletal- associated proteins [114].

Despite minimal sequence homology, the three-dimensional structures are remarkably conserved. As descripted above, the PH domains share the same PH superfold structure [115], [116] as the PTB domains; however, PH domains differ in their functional ability to bind membrane lipids. Essentially all PH domains bind phosphoinositides or inositol phosphates, with the exception of the PH domains from the Golgi-associated evectin proteins [116], [117]. Lipid- binding occurs through electrostatic interactions with the polarized, positively charged face of the PH domain, conferred by three positively charged loops of highly variable length and amino acid sequence [116]; however, the affinity and specificity of PH domains is highly variable [116], [118], depending on the presence or absence of variable charged side chains, which mediate hydrogen bonds with phosphate headgroups [119].

2.3. Roles of Adaptor Proteins in EGFR Signaling

EGFR activation and autophosphorylation provides multiple intracellular docking sites for adaptor proteins, such as Growth factor receptor-bound protein 2 (Grb2), Shc, Grb2- associated binder 1 (Gab1) and Gab2. These proteins play important roles in transducing EGFR signaling by recruiting downstream signaling molecules.

The cytosolic adaptor protein Grb2 is a classic example of SH2 domain-containing protein, comprised of a central SH2 domain flanked between an N-terminal SH3 (nSH3) domain and a C-terminal SH3 (cSH3) domain, giving it an overall modular architecture of nSH3–SH2– cSH3 [120]. Grb2 recognizes activated EGFR by virtue of its SH2 domain to bind to pTry sequences in the context of the consensus motif pYXN located within the cytoplasmic tails of a diverse array of receptors, including EGFR. Grb2 binds directly to the EGFR at phosphorylated Y1068, to a lesser extent at phosphorylated Y1086, and indirectly, via the adaptor molecule Shc [121] (Figure 3).

34 Functionally, Grb2 plays an essential role downstream of the EGFR by activating Ras through the recruitment of the RasGEF, son of sevenless (Sos1), via one of the SH3 domains of Grb2 and the C-terminal proline rich region of Sos1. The Grb2-Sos1 interaction is constitutive, independent of EGF stimulation; however, upon EGFR activation, Grab2 places it in close proximity to the plasma membrane, where it can mediate the exchange of GDP for GTP on Ras, activating membrane-bound Ras, leading to the initiation of the MAPK signaling cascade [121], promoting cellular proliferation. Furthermore, Grb2 can bind a PXXXR motif [122], which forms an SH3 domain binding site on other adaptor molecules, the insulin receptor substrate (IRS)-like Gab1 and Gab2, which can then recruit PI3K, via the Gab N-terminal PH domain, propagating the PI3K/Akt (also known as the PKB) pathway [102], [123], enhancing cell growth, proliferation, transcription and protein synthesis (Figure 3). Additionally, Grb2 can also negatively regulate EGFR signaling through receptor degradation via the recruitment of the E3 ubiquitin ligase Cbl, which interacts with the Grb2 SH3 domain, leading to receptor endocytosis [124].

As mentioned above, Grb2 can also couple to the EGFR indirectly through the cytosolic adaptor protein Shc, which comprises a C-terminal SH2 domain, a central glycine/proline-rich region and an adjacent PTB domain [125]. Shc binds the activated EGFR at phosphorylated Y1148 via is SH2 domain and phosphorylated Y1173 via is PTB domain [121], [125], [126], is tyrosine phosphorylated and recruits the Grb2-Sos complex [125] via the Grb2 SH2 domain, playing a major role in the Ras-MAPK signaling pathway as well as the Ras-ERKs-RSK pathway [127], regulating cell proliferation, survival, growth and motility [128] (Figure 3).

3. The Downstream of kinase (Dok) family of Adaptor Proteins

The mammalian Dok protein family comprises seven members (Dok-1–Dok-7), which share structural similarities characterized by the N-terminal PH and PTB domains followed by SH2 target motifs in the C-terminal moiety, indicating an adaptor function. Dok-1 was originally identified as a 62 kDa protein that binds with p120 rasGAP, a potent inhibitor of Ras, upon tyrosine phosphorylation by a variety of protein tyrosine kinases, including EGFR [129]. Based on their amino acid sequence, the Dok family can be divided into three subgroups: Dok-1, Dok-2

35 and Dok-3 form group A; Dok-4, Dok-5 and Dok-6 cluster into a second subgroup B; while Dok-7 is a more distantly related family member, forming a third subgroup [130] (Figure H). In addition to these members, a splice variant of Dok-4, Dok-4b, has been identified, containing a 39 amino acid insertion in the C-terminal region of the protein [131]. Dok adaptor proteins are primarily implicated in the negative regulation of tyrosine kinase signaling [130].

A Dok-1 PH PTB Y Y YY YYYY Y (481 aa)

Dok-2 PH PTB Y Y YYY Y (412 aa)

Dok-3 PH PTB Y Y Y Y (444 aa)

Dok-4 PH PTB YY Y YY (325 aa)

Dok-5 PH PTB Y Y Y (306 aa)

Dok-6 PH PTB Y Y Y YY (331 aa) Dok-7 PH PTB YY YY (504 aa)

Dok-1 B a a Dok-2 Dok-A subgroup (Dok-1/2/3) Dok-3 Dok-4 a Dok-5 Dok-B subgroup (Dok-4/5/6) a a Dok-6 Dok-7

Figure H: The Dok family proteins. (A) Schematic representation of mouse Dok proteins. Y denotes a tyrosine residue in the C-terminal region. (B) A phylogenetic tree of mouse Dok proteins. The Dok adapters cluster in three subgroups, namely Dok-A (Dok-1 ⁄ 2 ⁄ 3), Dok-B (Dok-4 ⁄ 5 ⁄ 6), and Dok-7. Adapted from: Mashima, R., et al., 2009 [130].

3.1. Structure

Structurally, the N-terminal region of the Dok family contains a tandem arrangement of PH and PTB domains. The unstructured C-terminal region shares little sequence homology amongst

36 Dok family members; however, it contains multiple tyrosine residues and proline-rich sequences, which may act as docking sites for SH2 and SH3 domain-containing proteins.

Protein tyrosine kinases that phosphorylate Dok proteins are localized to cellular membranes, as are many proteins involved in Dok-mediated signaling; therefore, the membranous localization of Dok proteins facilitate their activities in receptor signaling [132]. The PH domain plays an essential role in the localization or translocation to cellular membranes through its interaction with multiple phospholipids, including phosphatidylinositol 3,4-diphosphate

[PI(3,4)P2], phosphatidylinositol 4,5-bisphosphate [PI(4,5)P2], and phosphatidylinositol 3,4,5- trisphosphate [PI(3,4,5)P3] [130]. Functional studies have demonstrated that phosphoinositide recruitment of Dok-1 to the plasma membrane is essential for its negative regulation of MAPK activation [132].

The Dok-4 PTB domains are classified as IRS-like PTB domains [111], sharing 30-40% sequence identity with the IRS family PTB domains [133]. The crystal structure of the Dok-1 PTB domain was successfully resolved, demonstrating a b-sandwich composed of two nearly orthogonal, 7-stranded, antiparallel b-sheets, capped at one end by a C-terminal a-helix [134], typical to most PTB domains. The peptide-binding site in the Dok-1 PTB domain is characterized by an L-shaped surface groove formed by residues from the b5 strand and the C- terminal a-helix [134]. All Dok family PTB domains contain two conserved argenine residues, Arg207 and Arg222, which extend from the b5 and b6 strands, respectively, and are involved in recognizing and forming hydrogen bonds with a phosphotyrosine residue [134]. The important amino acid sequence in the ligand is usually initiated by a –3 aspargine residue, a –2 proline residue and a relative phosphorylated tyrosine, designated at position 0, forming the canonical PTB-binding NPXpY motif [111], [135]; however, systematic screening of all the Dok family members remains to be done, in order to determine their PTB binding motifs. Much of the function of the Dok family members is believed to occur through interactions with various binding partners, mediated by the PTB domain. Additionally, the PTB domain is also the site of homotypic and heterotypic oligomerization within the Dok family; however, the functional relevance of this oligomerization is unclear [130].

3.2. Expression

Members of the Dok family have both overlapping and tissue-specific expression, with members of the Dok-A subgroup (Dok-1/2/3) being preferentially expressed in immune cells, while members of the Dok-B subgroup (Dok-4/5/6) and Dok-7 are expressed in non- hematopoetic cells, with Dok-7 expression being limited to myocytes. Specifically, Dok-1 mRNA is expressed in B- and T-cells as well as in myeloid cells such as macrophages and neutrophils, with the exception of CD4-CD8- thymocytes and pro-B B220+CD43+IgM- lymphocytes [130], [136], [137]. Similarly, Dok-2 mRNA is expressed in T-cells and myeloid cells, with little or no expression in B-cells [136], [137]. By contrast, Dok-3 mRNA is preferentially expressed in myeloid cells and B cells, with little or no expression in T cells [137][138]. However, Dok-1 proteins have also been detected in non-hematopoietic cells, suggesting roles in non-immunological functions [129], [130], [139]–[142]. The expression of Dok-1/2/3 are thought to be absent from most epithelial cells. In contrast, Dok-4 mRNA is ubiquitously expressed, with high levels of expression in the kidney, intestine, lung, muscle, brain and heart [143], [144]. In situ hybridization showed that the Dok-4 transcript is expressed in the embryonic nervous system [143], and expression has also been detected in human T-cells [145]. The Dok-4b splice variant is expressed in epithelial cell lines, such as the adenocarcinoma cell line, Caco-2, the ductal breast cancer cell line, T47D and the cervical cancer cell line, HeLa [131]. Dok-5 and Dok-6 expression is mostly restricted to neural cells [143], [146]; however Dok-5 mRNA expression was also detected in human T-cells [145] and weak expression was found in muscles [144]. Modest expression of Dok-6 was also found in the kidney, spinal cord and testes [146]. Dok-7 expression is restricted to cardiac and skeletal muscle, and immunohistochemical analyses of skeletal muscle sections shows the distribution of Dok-7 to be restricted to the post-synaptic region of neuromuscular junctions [147].

Within the cell the, Dok family members are most often localized to the cytoplasm and to the membrane by their PH domain [130]; however both Dok-1 and Dok-4 possess a short leucine- rich motif, which acts as a functional nuclear export signal (NES) that can be recognized by a nuclear exporter such as Crm-1, allowing Dok-1 and Dok-4 to shuttle between the cytoplasm and

38 the nucleus [148], [149]. The function of Dok-1 and Dok-4 within the nucleus remains to be elucidated [148].

3.3.Function Functionally, the Dok family proteins have diverse roles in regulating signal transduction, primarily in the negative regulation of tyrosine kinase signaling. Commonly, the consequence of signaling involving the Dok proteins involves inhibition of cell growth and proliferation [130]. Depletion of the Dok-A subgroup (Dok-1/2/3) in mice leads to the development of aggressive, histiocytic sarcomas, a malignant proliferation of histiocyte cells with features of tissue-resident macrophages [150]. In vitro models have demonstrated roles for Dok family members as regulators of proliferation. Knockout of Dok-1 in mouse embryonic fibroblasts increased PDGF- induced proliferation and activation of the Ras/MAPK pathway and re-introduction of cytosolic Dok-1 expression restored proliferation and Ras/MAPK signaling to normal levels [132]. In the SKBR3 breast carcinoma cell line, overexpression of Dok-2 results in increased anoikis [151], a form of apoptosis, which is induced by cell detachment. A study in Dok-3-deficient mice found that Dok-3 modulates bone remodeling through the negative regulation of osteoclastogenesis by inhibiting activation of Syk and ERK in response to RANKL and M-CSF and through the positive regulation of osteoblastogenesis [152]. Commonly, expression of Dok-A family members results in the inhibition of the MAPK signaling cascade, downstream of tyrosine kinase receptors, achieved through interactions with various signaling partners [132], [141], [151], [152].

Less is known about the function of the Dok-B family. Ectopic overexpression of Dok-4 in 293 human embryonic kidney (HEK) and Caco-2 cells can inhibit the activity of Elk-1 [144], a ternary complex factor, which acts as a transcriptional activator, commonly activated downstream of ERK in response to growth factor stimulation; thus, supporting an inhibitory role for Dok-4 in epithelial cell proliferation. In T-cells, Dok-4 is a negative regulator of ERK phosphorylation, IL-2 promoter activity and both T-cell proliferation and activation [153]. The impact of Dok-4 on MAPK signaling appears to be dependent on cell context, as Dok-4 appears to positively regulate glial cell line-derived neurotrophic factor (GDNF)-dependent neurite outgrowth during neuronal development, through downstream activation of Rap1-ERK1/2 and

39 MAPK [154]. Additionally, Dok-4, Dok-5 and Dok-6 appear to positively regulate the MAPK pathway in neurons, downstream of the RTK, c-Ret, where they promote neurite outgrowth [143], [146].

Dok-7 is directly implicated in human disease, as mutations in Dok-7 disrupt the structure of the neuromuscular junction in patients with congenital myasthenic syndromes [155]. Dok-7 is essential for neuromuscular synaptogenesis, as mice deficient in Dok-7 are immobile and die shortly after birth [147]. Dok-7 directly interacts with the muscle-specific tyrosine-protein kinase receptor (MuSK), through its PTB domain. Because they spontaneously dimerize, Dok-7 molecules form clusters of acetylcholine receptors at the neuromuscular junction, which facilitate receptor activation and are necessary for proper neuromuscular synaptic transmission [147]. Thus, unlike other family members, Dok-7 acts as an intracellular RTK agonist.

3.4. Partners As the Dok proteins do not have any obvious domains for catalytic activity, it is likely that their binding partners and subcellular localization define their cellular functions.

The members of the Dok-A family all bind the lipid phosphatase, SHIP-1 [156]–[158]. This interaction occurs through the Dok PTB domain and requires the phosphorylation of not only SHIP-1 but also of the Dok members, suggesting the involvement of the SHIP-1 SH2 domain [156], in addition to the Dok-4 PTB domain, in the interaction between SHIP-1 and members of the Dok-A subgroup.

In addition to their interaction with SHIP-1, Dok-1/2/3 all interact with some form of the kinase Abelson (Abl) [129], [159], [160], although the mode of interaction differs among the members of the Dok-A family. The able kinase contains an SH3 domain, followed by an SH2 domain, a catalytic domain and an F- and G-actin bonding domain [160], [161]. Dok-1 and Dok- 3 can associate with Abl through their PTB domains, in a tyrosine kinase-dependent manner [162]. Interaction between Dok-1 and the oncogenic BCR-Abl fusion protein was found to require tyrosine phosphorylation of Dok-1 and the SH2 domain of BCR-Abl [163]. Dok-2

40 constitutively binds the SH3 domain of Abl [151], through a proline-rich PMMP motif in its C- terminal [160].

Csk is a cytoplasmic tyrosine kinase consisting of an SH3, an SH2, and a kinase domain [164], that interacts with all the members of the Dok-A family. Dok-1 recruits Csk to the plasma membrane upon PDGF stimulation, through its Y449, decreasing Src kinase activity and PDGF-induced mitogenesis [141]. Downstream of EGFR activation, phosphorylation of Y402 on Dok-2 facilitates the recruitment of Csk, down-regulating Src activity and both MAPK and Akt/protein kinase B (PKB) activity [165]. In B-cells, tyrosine phosphorylation of Dok-3 in response to immunoreceptor-mediated cellular activation induces its binding to Csk [138].

Both Dok-1 and Dok-2 interact with the Ras GTPase activating protein (GAP), p120RasGAP [129], [166], with the result of Dok-1 inhibiting the activation of Ras and ERK, attenuating MAPK signaling [130] and in the case of Dok-2, inhibiting T-cell development [167]. Similarly, both Dok-1 and Dok-2 were found to interact with two known tyrosine phosphorylation sites at Y1086 and Y1148 in EGFR via their PTB domains [168], [169], acting as negative regulators of EGFR signaling; whereas tyrosine phosphorylation sites on Dok-3 allow it to bind with Grb2 via its SH2 domain, sequestering Grb2 from Shc and inhibiting Ras-ERK sgnaling downstream of RTKs [170]. Additionally, both Dok-1 and Dok-2 can interact with the adaptor protein Nck, downstream of insulin receptor (IR) activation and Dok phosporylation, inhibiting insulin- stimulated activation of Ras and Akt [171] and enhancing insulin-induced cell migration [172], respectively.

The most well-characterized binding partner of the Dok-B family is the receptor tyrosine kinase for members of the GDNF family of extracellular signalling molecules, Ret. Ret binds Dok-1, suppressing GDNF-induced Ras/ERK activation, through its association with p120RasGAP, and enhancing GDNF-induced activation of the c-Jun amino-terminal kinase (JNK) and c-Jun activation, through its association with Nck [173]. In neuronal cells, Ret also interacts with Dok-4, Dok-5 and Dok-6 to enhance GDNF-mediated neurite outgrowth [143], [146]. In T-cells, the Dok-4 PTB domain binds the lipid phosphatase SHIP-1 [174]. In epithelial

41 cells, insulin receptor-phosphorylated Dok-4 is found to bind RasGAP, the adaptor protein Crk, and the tyrosine kinases Src and Fyn, with the effect of activating MAPK signaling [133]. It was also found that Dok-4 directly binds to and inhibits the transcription factor Elk-4, thereby inhibiting EGF-induced epithelial cell proliferation [149]. Due to their restricted expression, most of the Dok-5 and Dok-6 binding partners are neuron-specific. Dok-5 directly interacts with tropomyosin-related kinase (Trk) receptors, TrkB and TrkC, high affinity receptors for neurotrophin growth factors, through a kinase-dependent PTB domain-mediated interaction, resulting in neurotrophin-induced MAPK signal pathway activation [175]. Dok-6 is also found to bind TrkC via its PTB domain in a kinase-dependent manner, promoting neurite outgrowth in primary cortical neurons [176].

As mentioned in the previous section, the only known binding partner of Dok-7 is the MuSK receptor, binding to which is necessary for proper neuromuscular synaptogenesis [147].

3.5. Dok-4

Dok-4 is the only member of the Dok family that is ubiquitously expressed; it is widely expressed at low levels in multiple tissues and is enriched in neural, endothelial and epithelial tissues, yet despite its wide expression pattern, the functions of Dok-4 and the Dok-4b splice variant remain poorly defined. The fact that it is the only member of the Dok family to be highly expressed in cells and tissues of epithelial origin, suggests that Dok-4 may have roles distinct from the other members of the Dok family.

Dok-4 is constitutively membrane localized in mammalian cells and yeast, and can be tyrosine phosphorylated by multiple kinases including Src, Fyn, Jak2 and Ret; membrane- localization as well as phosphorylation by Fyn and Ret require the Dok-4 PTB domain [144]. Dok-4 was originally identified, along with Dok-5, as a binding partner and substrate of the Ret receptor [143]; however, for the purpose of this project, we are not studying Dok-4 in this context, as Ret expression is absent in mature epithelium. Previous studies, using a yeast two- hybrid system found that Dok-4 does not directly interact with the EGFR [143]; however, in this

42 study, we are investigating Dok-4 interactions downstream of the EGFR, as both Dok-4 and Dok-4b appear to inhibit EGF-induced Elk-1 activation in Caco-2 adenocarcinoma cells [131], implicating Dok-4 as a negative regulator of epithelial cell proliferation. Additionally, in Caco-2 cells, Dok-4 expression is highly upregulated following contact-dependent cellular differentiation [144], suggesting that Dok-4 contributes to maintain quiescence in differentiated epithelial cells. This was confirmed in Maden-Darby canine kidney (MDCK) renal tubular epithelial cells, where siRNA-mediated knockdown of endogenous Dok-4 enhanced cellular proliferation, associated with an upregulation of cyclin D1 mRNA and a downregulation of the cyclin dependent kinase inhibitor p21Cip1/Waf1, both basally and in response to EGF stimulation [149]. Taken together, these findings support the notion that Dok-4 inhibits cell cycle progression through modulation of signaling upstream of cyclin D1. However, the exact mechanisms for this action remain unknown.

The key to understanding the action of the Dok-4 adaptor protein, is to identify its binding partners. In addition to Elk-4, we have identified through yeast two-hybrid screening of a mouse cDNA library, the Rac-specific Rho-GTPase activating proteins (GAPs) α2-chimaerin (Chn1) and β2-chimaerin (Chn2), as two novel direct binding partners for Dok-4 in epithelial cells. Elk-4 and the Chn1 and Chn2 proteins are all highly upregulated in the reperfused kidney, following ischemic insult, suggesting that the interaction of Dok-4 with these two proteins may be implicated in a negative feedback mechanism to growth factor-induced proliferation following renal IRI. Continued investigation is needed to understand the functional impact of Dok-4, a unique epithelial Dok family adaptor protein.

4. Chn family of Rac-GAPs The chimaerin (Chn) family of proteins comprise five known members: a1-, a2-, b1-, b2- and b3-chimaerin. Chimaerins are the only known RhoGAPs to bind phorbol ester (PE) tumour promoters and the lipid second messenger diacylglycerol (DAG) and show specific GAP activity towards the small GTPase Rac. Studies have established that chimaerins are regulated by tyrosine kinase and G-protein-coupled receptors (GPCRs) via PLC activation and DAG generation to promote Rac inactivation. The finding that chimaerins, and some other proteins, are

43 receptors for DAG changed the prevalent view that PKC isoenzymes are the only cellular molecules regulated by DAG. In addition, recent studies have begun to decipher the critical roles of chimaerins in the central nervous system, development and tumour progression.

4.1. Structure

Five isoforms of chimaerin have currently been identified, which are products of CHN1 gene (a1- and a2-chimaerins), mapped to chromosome locus 2q31-32.1 and CHN2 gene (b1-, b2- and b3-chimaerins), mapped to chromosome locus 7p15.3 [177], [178] (Figure I). Although these gene products were initially assumed to be generated by alternative splicing, bioinformatics analysis of the CHN2 gene revealed that b1-, b2- and b3-chimaerins are the products of alternative transcription start sites (TSSs) in different promoter regions of the CHN2 gene [178].

b1-chimaerin C1 GAP

CHN2 b2-chimaerin P SH2 C1 GAP

b3-chimaerin SH2 C1 GAP

a1-chimaerin C1 GAP CHN1 a2-chimaerin P SH2 C1 GAP

Figure I: The Chimaerin Family Proteins. Schematic representation of the domains structures among human chimaerin proteins. Domain abbreviations: P, proline-rich domain; SH2, Src homology 2 domain; C1, C1 domain; GAP, GTPase activating protein domain.

Chimaerins were originally named for the fact that they resemble a ‘chimaera’ between the C1 domain of PKC isoenzymes and the GAP domain of breakpoint cluster region (BCR), involved in Philadelphia chromosome translocation [77]. All five chimaerins have a C-terminal

44 GAP domain and a single PKC-like C1 domain. The α2-, β2- and β3-chimaerins have an additional N-terminal SH2 domain.

The chimaerin C1 domains are small cysteine-rich structural units of about 50 amino acids, originally defined as lipid-binding modules in PKC isoenzymes. C1 domains can be classified as “typical”, DAG/PE-binding or “atypical”, non-DAG/PE-binding [179]. The chimaerins contain a single [179] “typical” C1 domain, which binds the lipid second messenger DAG and DAG- mimetics such as PEs and is critical for governing association to membranes. The C1 domain in chimaerins is about 40% homologous with C1 domains in PKC isoenzymes and possesses the structural requirements for phorbol ester binding, including the motif

HX12CX2CX13/14CX2CX4HX2CX7C (where H is histidine, C is cysteine, X is any other amino acid), characteristic of other typical C1 domains [77]. Modelling studies determined the chimaerin C1 domain to have a globular fold structure with a PE binding groove on top and to contain characteristic hydrophobic amino acids that participate in membrane insertion [180]. As chimaerin genes alternative TSS/alternative splicing occur upstream of the C1 domain, all isoforms from each gene have identical C1 domains [178], [181]–[183]. Functionally, the C1 domain is crucial for the positional regulation of the chimaerin proteins, as chimaerins redistribute to membranes in response to PE/DAG activation. A series of site-directed mutagenesis and deletional analyses confirmed in β2-chimaerin, that the C1 domain is essential for its ability to bind PEs and to translocate to membranes [180], [184]. Mutation of an essential Cys246 to alanine in β2-chimaerin, which disrupts the overall structure of the C1 domain, impairs the ability of the protein to relocalize in response to phorbol myristate acetate (PMA) or other PE analogues [180]. This analogy with PKC isoenzymes led to the hypothesis that chimaerins could be effectors for DAG generated in response to receptor activation.

The chimaerin GAP domain is composed of about 150 amino acid residues and it is highly homologous with the BCR GAP domain [185], [186]. The GAP domains of α-and β-chimaerins share approximately 77% identity [183]. β2-chimaerin has been shown to preferentially accelerate the hydrolysis of GTP from Rac with a roughly 50-fold higher rate than from Cdc42, while no activity for Rho was detected [180], [187]. The RacGAP activity of chimaerins is

45 sensitive to the lipid environment, specifically, it is stimulated by the presence of PEs/DAG, PS (phosphatidylserine) and PA (phosphatidic acid) [188].

The α2- and β2- and β3-chimaerins all have N-terminal SH2 domains that contain a glutamic acid residue at the start of the domain instead of the tryptophan residue commonly found in other SH2 domains; however, the conserved arginine residues and the phosphotyrosine- binding sequence are preserved [77]. Little is known about the biological roles of these domains in chimaerins. While studies have shown that the SH2 domain of RasGAP binds to various growth factor receptors [189], it is not known whether the SH2 domains of α2-, β2- and β3- chimaerins have similar properties. Interestingly, studies have demonstrated that the α2- chimaerin SH2 domain interacts with other proteins, such as the microtubule-associated protein collapsin-response mediator protein-2 (Crmp2), and that this interaction is required for neuritogenesis [190], [191].

Much of our understanding of the regulation and function of chimaerins has been achieved with the resolution of the three-dimensional structure of β2-chimaerin, the first structure of a full-length protein with a PE-responsive C1 domain to have been solved [77]. Data from crystallographic analysis revealed autoinhibitory mechanism, also found in α2-chimaerin [192] (Figure J), whereby the C1 domain forms extensive hydrophobic contacts with the GAP domain and the SH2 domain. The N-terminus of β2-chimaerin protrudes into the active site of the RacGAP domain, sterically blocking Rac binding. The DAG-binding site in the C1 domain is buried by contacts with the N-terminus, the SH2 domain, the RacGAP domain and the SH2-C1 linker region. These structural characteristics imply that the C1 domain in β2-chimaerin is inaccessible to DAG or PE when the protein is in this conformation, suggesting the need for a conformational change that exposes the C1 domain in order to bind DAG. Lipid binding to the C1 domain requires the co-operative dissociation of these intramolecular interactions, allowing the N-terminus to move out of the active site and enzyme activation [193]. These structural observations were confirmed by mutagenesis studies, in which a series of β2-chimaerin mutants that destabilize the inactive conformation were generated [193]. Remarkably, when these mutants were tested for their ability to translocate from the cytosolic fraction to membranes in

46 response to PMA, a PKC activator, it was found that they respond much more readily to PMA than wild-type β2-chimaerin, particularly the mutant I130A-β2-chimaerin, which has an approximately 100-fold lower EC50 for the phorbol ester than the wild-type β2-chimaerin [193]. The RacGAP activity of I130A-β2-chimaerin is also enhanced; whereas wild-type β2-chimaerin reduces Rac-GTP levels by about 25% in COS-1 cells, the mutant I130A reduces Rac-GTP levels by about 80% [193].

EGFR

PM PIP2 DAG

SH2 Rac-GTP PLCɣ P P Nck C1 RacGAP Rac-GEF P

SFK Rac-GDP

P SH2 C1

RacGAP

SH2 C1

RacGAP α2/β2-chimaerin

Figure J: Model for the EGFR-Mediated Regulation of Chimaerins. α2/β2-chimaerin are in a closed autoinhibited conformation in the cytosol. EGFR stimulation leads to Rac activation and possible phosphorylation of the chimaerins causes a conformational change to the open active conformation. The PLCγ branch of EGFR leads to DAG generation, and translocation of chimaerins, where they are stabilized at the membrane through their interaction with DAG via their C1 domain and with adapter proteins (Nck) via a proline rich N-terminal region and are able to inactivate Rac. Adapted from: Wang, H.B. et.al., 2006 [194].

Based on these findings, a model for α2- and β2-chimaerin activation has been proposed. The inactive proteins reside in the cytoplasm; when cells are stimulated with PEs or growth

47 factors trigger generation of DAG, a conformational change occurs and the C1 domain is exposed, an event that facilitates ligand binding, membrane insertion, association with Rac and activation of chimaerin RacGAP activity. Although this model explains how the enzyme activity of chimaerin is activated, it does not tell us how the conformational changes occur. The prediction is that other events, such as post-translational modifications (phosphorylation) or interactions with yet unidentified proteins, must be required to facilitate the transition to the open conformation.

Tyr phosphorylation in PKCθ leads to a conformational change that exposes the C1 domains for membrane interactions [195] and evidence suggests that Tyr phosphorylation of α2- and β2-chimaerin modulates their activation and facilitates their redistribution to the membrane [196], [197]. DAG-dependent protein-protein interactions involving a proline rich motif in the N- terminal region of α2- and β2-chimaerins and acidic clusters in the SH3 domain of the adaptor protein Nck1, are necessary for the translocation of chimaerin and activation of its GAP activity downstream of the EGFR [196]. It is unclear what role the SH2 domain plays in chimaerin activation. It is conceivable that SH2 domain interaction with tyrosine phosphorylated proteins, including receptors, contributes to the process of chimaerin activation and probably co-operates with the C1 domain for membrane association.

4.2. Expression

Studies show that α1-chimaerin is primarily expressed in brain [185]; rat α1-chimaerin mRNA is restricted to neurons, with the highest levels found in hippocampal pyramidal cells, granule cells of the dentate gyrus and cortical neurons; in cerebellum α1-chimaerin mRNA can only be detected in Purkinje neurons [198]. β1-chimaerin is mainly expressed in testis [183]. α2- chimaerin is highly expressed in brain, particularly in neuronal perikarya, dendrites and axons [190], while β2-chimaerin is highly expressed in cerebellum and is mainly expressed in granule cells [181]; however, both α2- and β2- chimaerin mRNA can be detected in a wide range of cultured cell lines [77]. Additionally, β2-chimaerin has been also found in T-lymphocytes [199]. Within the cell, PEs and DAG analogues induce the intracellular translocation of chimaerins from cytosolic (soluble) to membrane (particulate) fractions [180], [184], [200], [201].

48 4.3. Function

Compared to our understanding of the biochemical properties of chimaerins, relatively little is known about their biological functions. Acting as inhibitors of Rac activity, an important modulator of the cytoskeleton, chimaerins have been implicated in regulating development and cellular morphology and can act as negative regulators of cellular proliferation and migration.

Since chimaerins are highly expressed in brain, it is expected that they should play roles in regulating Rac-mediated responses in the central nervous system (CNS). In cultured hippocampal neurons, α1-chimaerin is capable of modulating dendritic spine density [202] through binding to synaptic N-methyl-D-aspartate (NMDA) receptor NR2A subunits and locally inactivating Rac1 [203]. Both DAG- binding and RacGAP activities of α1-chimaerin were required for the induction of the pruning of dendritic spines and branches [202], [203]. The SH2 domain containing α2-chimaerin is also implicated in regulating hippocampal pruning. Investigation into the functional role of the α2-chimaerin SH2 domain in neurons has found transient expression of α2- chimaerin, but not α1-chimaerin, in N1E-115 neuroblastoma cells leads to neurite formation, independent of nerve growth factor (NGF) stimulation [190]. In contrast, transfection with α2-chimaerin mutated in the SH2 domain (N94H) inhibits neurite formation in NGF-stimulated PC12 cells, indicating a role for α2-chimaerin in neuritogenesis for which its SH2 domain is indispensable [190].

Chimaerins are highly expressed in the rat embryonic nervous system [181], [190] suggesting a potential role during development. In zebrafish, the CHN1 gene product expresses the three modules present in α2- and β2-chimaerins (SH2, C1 and GAP domains), and it is highly homologous with the mammalian α2-chimaerin isoform, biochemically possessing both RacGAP activity and PE/DAG binding capability [204]. The chn1 transcript is detected in zebrafish embryos as of the eight-cell-stage and is widely distributed during the cleavage, blastula and gastrula periods, while during the segmentation period, the expression of chn1 is restricted to the neural tissue, and, by the pharyngula period, the protein is highly expressed in brain [204]. At the larval stages, chn1 is also present in liver, gut, pancreas and pharyngeal structures, coinciding with the expression pattern of its putative target, zebrafish Rac1 (rac1) [204].Chn1 morpholino

49 knockdown embryos exhibit severe abnormalities, including the development of round somites, lack of yolk extension, and a kinked posterior notochord; these morphants show Rac hyperactivation and progress faster through epiboly, leading to tailbud-stage embryos that have a narrow axis and an enlarged tailbud with expanded bmp4 and shh expression [204]. Mutational studies demonstrated that the lack of chn1 RacGAP activity in the yolk syncytial layer is the main cause of the defects in morphogenetic movements [204]. These studies reveal an important role for chn1 in early development, but also implicate Rac as a key regulator of morphogenetic movements during zebrafish epiboly. In mouse embryonic CNS development, α-chimaerin is required to establish the midline barrier for proper corticospinal axon guidance [205]. A study of cytogenesis in epithelial MDCK cells implicated β2-chimaerin-induced suppression of Rac1 at the apical membrane as being essential for the maintenance of cyst structure [206]. Considering that many sporadic cancers arise from such polarized tissues, disruption of the regulation of polarized Rac1 activity might be a common step in the initiation of cancerous transformation in cells. The well-established role for Rac and its effectors on cell proliferation and motility [207] also suggest potential roles for chimaerins in human cancer development and progression.

It is now widely accepted that Rac and other Rho GTPases play important roles in regulating cell morphology and movement, invasion, metastasis, proliferation and malignant transformation, which are all crucial events in cancer development and progression [75], [208]. Both Rac1 and its spliced variant Rac1b are highly expressed and hyperactive in human cancers [208]–[210]. It has been hypothesized that chimaerins or other RacGAPs possess tumour- suppressor capabilities, and accumulating evidence supports this hypothesis. The potential link between chimaerins and cancer was first investigated in astrocytoma [211]. Human astrocytomas are the most common primary CNS tumours, but the molecular mechanisms leading to the malignant transformation from low- to high-grade tumours are not well understood; however, β2-chimaerin is found to be differentially expressed in brain tumours, with high expression levels detected in normal brain and low-grade astrocytoma and low expression levels detected in malignant high-grade gliomas [211]. These findings support a role for β2-chimaerin as a tumour- suppressor gene and implicate that dysregulation of Rac activity may play a role in the progression of human brain cancers. Similarly, the transcript levels of β2- chimaerin in human breast cancer cells are significantly lower than in immortalized normal breast cells and β2-

50 chimaerin mRNA levels in human breast cancer tissues are also significantly lower than those in paired normal breast tissues from the same patients [212], supporting a role for chimaerins in tumour suppression. Stable transfectants in mouse mammary carcinoma cells expressing the β2- chimaerin GAP domain have reduced proliferation rates and invasiveness capability in vivo, as ectopic expression of β2-chimaerin induces cell-cycle arrest in G0 /G1 and inhibits the proliferation of MCF-7 breast cancer cells in a RacGAP-dependent manner [213]. Reduction in Rac-GTP levels by expression of the β-GAP chimaerin domain in MCF-7 cells correlates with decreased expression of cyclin D1, reduced retinoblastoma family protein (pRb) phosphorylation levels and inhibition of cell-cycle G1/S transition [213]. Recent studies have pointed to a crucial role for Rac as a mediator of growth factor responses in breast cancer cells [212], [214]. EGF and HRG (heregulin β1) cause significant elevations in Rac- GTP levels in breast cancer MCF-7 [212] and T47D [214] cells and promote breast cancer cell migration and proliferation in a Rac- dependent manner. β2-chimaerin inhibits HRG-induced breast cancer cell migration and proliferation through the inactivation of Rac [214], pointing to a role for β2-chimaerin in regulating growth factor-mediated responses that depend on Rac.

In addition to the anti-proliferative and anti-migration actions of β2-chimaerin on cancer cells, similar effects are seen in vascular smooth muscle cells, where ectopic expression of β2- chimaerin inhibits PDGF-induced Rac activation, thus impairing smooth muscle cell migration and proliferation [215], pointing to a potential role for chimaerins in human atherogenesis. In T- cells, expression of β2-chimaerin inhibits levels of active Rac, an effect that depends on its RacGAP activity and requires DAG generation [216]. Interestingly, while β2-chimaerin reduces cell adhesion, it enhances CXCL12-dependent migration through receptor-dependent DAG production [216]. Additionally, a C1 domain mutant (F215G) inhibited the effect of wild-type β2-chimaerin on PMA-induced integrin-dependent adhesion, and prevented the inhibitory effect of wild-type β2-chimaerin on integrin-dependent adhesion following CXCL12 stimulation [216], demonstrating an absolute requirement of an intact C1domain for inhibition of β2-chimaerin on cell adhesion.

4.4. Partners

51 Several studies have provided evidence of chimaerin-interacting proteins, suggesting that they may be part of multiprotein complexes [77]. Since chimaerins show a perinuclear localization and co-localize with a Golgi network marker in response to phorbol ester treatment [180], it is of no surprise that Tmp21-I (p23), a transmembrane protein localized in the cis-Golgi network and involved in intracellular trafficking, was identified as a chimaerin-binding protein in a yeast-two hybrid screening using β2-chimaerin as a bait [201]. The association of β2-chimaerin with Tmp21-I is promoted by phorbol esters in a PKC-independent manner and deletional analysis determined that chimaerins require an intact C1 domain for their interaction with Tmp21-I [201]. These findings have important mechanistic implications, as they reveal a dual role for the chimaerin C1 domain, both as a module for lipid recognition and for protein–protein interaction.

The p35 activator of cyclin-dependent kinase 5 (Cdk5) is an α-chimaerin-interacting protein and α-chimaerin, p35 and Cdk5 have been co-immunoprecipitated from HeLa cells, with both the Cdk5 kinase activity and the α-chimaerin GAP activity being retained in the protein complex [217]. Since Cdk5 in association with its neuronal activator p35 regulates neurocytoskeletal dynamics, it has been proposed that the association of α-chimaerin with Cdk5– p35 may be required for their co-ordinated involvement in the remodelling of neuronal actin filaments [217]. In dorsal root ganglion neurons of the CNS, active α2-chimaerin interacts with Cdk5–p35 through its GAP domain and with the Cdk5 target Crmp2 through its SH2 domain [218], suggesting that the α-chimaerin RacGAP activity plays an important role in regulating the dynamics of neurite outgrowth. Additionally, as previously mentioned, α1-chimaerin modulates dendritic spine density through binding to synaptic N-methyl-D-aspartate (NMDA) receptor NR2A subunits and locally inactivating Rac1 [203]. The identification of chimaerin-interacting proteins in neuronal models fits with the observation that chimaerins are highly expressed in the nervous system and with the proposed roles they may play in regulating functions in the CNS.

Also, as mentioned above, both α2- and β2-chimaerins interact with the Nck1 adaptor protein, to facilitate the translocation of chimaerin downstream of the EGFR [196]. β2-chimaerin is also found to interact with diacylglycerol kinase (DGK)g at the plasma membrane in COS7

52 cells, downstream of EGFR activation, enhancing EGF-dependent translocation of β2-chimaerin to the plasma membrane and markedly augmenting the EGF-dependent GAP activity of β2- chimaerin through its catalytic action [219]; suggesting that DGKg is a regulator of β2- chimaerin. In the CNS, α2-chimaerin binds to EphA4 receptors and mediates EphA4-dependent corticospinal axon guidance, while in HeLa cells, β2-chimaerin binds to EphA2 and EphA4 receptors and inactivates Rac1 in response to ephrinA1 stimulation, regulating cell migration [220]. A greater understanding of the interaction between chimaerins and their binding partners will help us glean insight into their functional role in epithelial cells.

53 Rationale and Objectives

The purpose of this study is to investigate the structure and functional role of the interaction between Dok-4 and the and β2-chimaerin in renal epithelial cells, downstream of the EGFR. Rac signaling has been implicated in many epithelial processes, in both normal and injured cells. Importantly, Rac regulates cellular proliferation, migration and polarization. Understanding the molecular basis underlying these events will be essential for pharmacologically targeting these processes during pathological states, such as cancer and ischemia reperfusion injury (IRI). This study aims to:

i) Identify the binding sites involved in the Dok-4/β2-chimaerin interaction. ii) Clarify the functional role of tyrosine phosphorylation in this interaction. iii) Characterize the functional role of the Dok-4/β2-chimaerin interaction on downstream Rac1-signaling.

CHAPTER 2: METHODOLOGY AND RESEARCH FINDINGS

55 METHODOLOGY

Yeast two-hybrid (Y2H) screening. Y2H screening was performed by Hybrigenetics (Paris, France) using full-length mouse Dok-4 C-terminally fused in-frame to LexA DNA-binding domain in vector pB27 as bait, as recently described [174]. To allow detection of phosphotyrosine-dependent interactions, a yeast strain expressing mammalian Lyn tyrosine kinase was used. After 75 million and 90 million possible interactions were screened with and without co-expressed Lyn tyrosine kinase, respectively, against a library of random-primed mouse kidney cDNAs fused to the Gal-4 activation domain.

Plasmids and cDNAs. The Dok-4 constructs were generated in the following vectors: pcDNA3.1(-) (Invitrogen), pcDNA3.1(-) Myc/His version A (Invitrogen), pEGFP-N1 (Clontech), pEGFP-C1 (Clontech), pmCherry-C1 (Clontech) and pGEX2T (GE Healthcare). The following Dok-4 constructs were described previously [131], [221]: pcDNA Dok-4-Myc/His (aa 1-325), pcDNA-Dok-4-Myc/His PTB (aa 100-246), pcDNA-Dok-4-Myc/His PTB G207A (aa 100-246, G207A), pcDNA Dok-4-Myc/His ΔPH (aa 100-325), and pcDNA Dok-4-Myc/His Myr ΔPH (aa 100-325). The additional Dok-4 construct, pGEX2T-Dok-4 PTB (aa 100–246), was generated by PCR cloning from the original Dok-4 expression vectors described above. The Dok-4-Venus construct was obtained by fusing a Venus aa 158–238 sequence to the C-terminus of Dok-4 in pCDNA 3.1., using a flexible (GGGGS)x2 linker.

The human b2-chimaerin (Chn2) (accession #BC 112155.1, in pCR4-TOPO) was obtained from ImaGenes. For expression in mammalian cells, Chn2 was sub-cloned into pcDNA3.1. The Chn2 NT (aa 2-182) construct was created by truncation at the internal XhoI and HindIII (both from New England Biolabs) sites of Chn2, and sub-cloned in frame into p-EGFP-C1, while full-length EGFP-Chn2 was obtained by sub-cloning the C-terminal fragment in frame into pEGFP-C1 Chn2 NT. The Chn2 Y153F mutant was obtained by sub-cloning a double stranded synthetic DNA fragment (Integrated DNA Technologies) with substitution of aa 153, in frame into pcDNA3.1. Chn2, using a Gibson assembly kit (New England Biolabs). The pEGFP-Chn2 Y153F mutant was obtained by sub-cloning pcDNA3.1. Chn2 Y153F in frame into pEGFP-C1. The Chn2 Y153A mutant was generated by overlap PCR with the sense oligonucleotide

56 AACCCCATCGCTGAACACATTGAATATGCC and the antisense oligonucleotide AATGTGTTCAGCGATGGGGTTAGTTGTCAT. The BGHR antisense and CMV sense oligonucleotides (Invitrogen) derived from pcDNA3.1. Chn2 were used to complete the reactions, and a second PCR was performed using the T7 sense and BGHR antisense oligonucleotides. The final PCR product was used for an HindII/XhoI insertion into pcDNA3.1. (-) (Invitrogen). The Venus-Chn2 and Venus-Chn2 Y153A constructs were obtained by fusing a Venus (aa 1–157) sequence to the N-terminal end of Chn2 and Chn2 Y153A in pCDNA3.1., using a flexible (GGGGS)x2 linker. The Chn2I130A mutant was obtained by sub-cloning a double stranded synthetic DNA fragment with substitution of aa 130 (Integrated DNA Technologies), into pEGFP-Chn2 and N-terminal tagged Venus-Chn2 I130A was created as described above.

To make a2-chimaerin (Chn1) the mouse Myc/Flag-a1-chimaerin (accession # NM_175752), was purchased from Origene and a2-chimaerin was created by replacing the N-terminal portion of a1-chimaerin with a double stranded synthetic DNA fragment (Integrated DNA Technologies) containing the N-terminal SH2 domain of a2-chimaerin, in frame using a Gibson assembly kit (New England Biolabs). The Chn1 Y143A mutant was obtained by sub-cloning a double stranded synthetic DNA fragment with substitution of aa 143 (Integrated DNA Technologies), in frame into pcDNA3.1. Chn1, using a Gibson assembly kit (New England Biolabs). The expression vector for activated (Y528F) Fyn was also described previously [221]. The pCDNA3 c-Src SH1 kinase catalytic domain (aa 250-536)-HA (Addgene) was graciously donated by Dr. R. Lefkowitz and the pXM-EGFR was graciously provided by Dr Morag Park. The GFP-tagged CA Rac1L61 and DN Rac1N17 were generously provided by Dr. N. Lamarche- Vane. Expression plasmids were sequenced by Genome Québec Innovation Centre.

Chemicals. Epidermal growth factor was purchased from BioSource and cells were treated with a concentration of 100 ng/mL. Isopropyl-1-thio-β-D-galactopyranoside and dithiothreitol were purchased from Sigma-Aldrich.

Cells and transfection. 293 HEK cells were cultured in DMEM-high glucose containing pyridoxine-HCL and sodium pyruvate (Invitrogen) with 10% FBS (Invitrogen). They were

57 incubated at 37°C, 5% CO2 and 90% humidity. Plasmid transfections were performed using Lipofectamine 2000 reagent (Invitrogen).

Antibodies and immunoblotting. Antibodies for GST, GFP and Myc tag (9E10) were purchased from Santa Cruz; anti-α-tubulin was from Sigma; anti-phosphotyrosine (4G10) was from Millipore; anti-Chn2 was from New England Biolabs and anti-Flag tag was purchased from Genscript. Dok-4 antiserum was generated in our lab as previously described [221]. Horseradish peroxidase-coupled secondary antibodies were from Jackson Immunoresearch and Rockland Immunochemicals, while fluorescent-labeled secondary antibodies were from LiCor. Lysates were obtained in 1X Laemmli gel-loading buffer or in standard IP buffer containing 1% Nonidet P-40 and protease inhibitors, as previously described [144]. Lysates were denatured by boiling for 5 minutes at 105°C and resolved on an SDS-polyacrylamide gel or TGX Stain-Free™ FastCast™ acrylamide gel (BioRad). StainFree™ gels were activated for 45 seconds, and imaged for total protein content, using the BioRad Touch Imaging system and transferred to PVDF or nitrocellulose membranes using the Trans-Blot® Turbo™ system (BioRad). Blots were revealed with ECL (Thermo Scientific) or ECL Prime (GE Healthcare) reagents, using x-ray film or the BioRad Touch Imager and ImageLab software (BioRad). Fluorescent-labeled secondary antibodies were revealed using the LiCor machine and Image Studio Lite Ver 5.2 software (LiCor).

Immunoprecipitation and Immunoblotting. 293 HEK cells were transiently transfected with Lipofectamine 2000 (Invitrogen). Forty-eight hours post-transfection, the cells were rinsed with ice-cold PBS and lysed on ice in standard IP buffer containing 1% Nonidet P-40 and protease inhibitors. IP was performed using 1-2 ug of affinity purified antibody or 5 uL of Dok-4 antiserum or 10 uL normal pre-immune rabbit serum followed by pull-down with 15-20 uL of protein A agarose beads (Santa Cruz). For IP with mouse antibodies, the protein A agarose beads were coated with 5 ug affinipure rabbit anti-mouse IgG (H+L) (Jackson ImmunoResearch). Immunoprecipitates were eluted in 1X Laemmli buffer, boiled for 5 minutes at 105°C and resolved on an SDS-polyacrylamide gel or TGX Stain-Free™ FastCast™ acrylamide gel (BioRad). StainFree™ gels and immunoblotted as described above

58 GST Fusion Proteins and in Vitro Binding. To create GST Dok-4 PTB, the pGEX2T-Dok-4 PTB (aa 100–246) fusion proteins were induced in BL21 Escherichia coli by the addition of 1 mM isopropyl-1-thio-β-D-galactopyranoside for 4 h at 30 °C. After lysis and sonication in IP buffer containing protease and phosphatase inhibitors, GST or GST fusion protein was coupled to glutathione-agarose beads (Sigma) in the presence of 5 mM dithiothreitol, for 2 h at 4 °C. Protein concentration was estimated by resolving aliquots on SDS-polyacrylamide gels and staining with Coomassie Blue or run on TGX Stain-Free™ FastCast™ acrylamide gels (BioRad) and activated and imaged for total protein content as described above. An in vitro binding assay was performed by incubating lysates of transiently transfected 293 cells with 20 uL of GST- or GST fusion protein-coupled beads for 1.5 h at 4 °C in the presence of 5 mM dithiothreitol. After five washes, proteins were eluted in 1X Laemmli buffer, boiled for 5 minutes at 105 °C, and loaded onto SDS-polyacrylamide gels. Immunoblots were carried out as described above.

Immunofluorescence Microscopy. 293 HEK cells plated on fibronectin-coated coverslips were transfected with expression plasmids using Lipofectamine 2000 reagent (Invitrogen) 24 h after plating and coverslips were processed 24 h after transfection. Coverslips were washed in PBS, fixed in 4% paraformaldehyde (Thermo Scientific) and permeabilized in 0.1% Triton X-100 (Sigma-Aldrich). Cells were then blocked using 3% BSA and incubated in mouse anti-Myc primary antibody (Santa Cruz) for 1.5 h at a dilution of 1:100 in 3% BSA (BioShop). Rhodamine red-anti-mouse secondary antibody (Jackson Immunoresearch) was diluted at a concentration of 1:1000 in 3% BSA. Following three washes, cells were stained with the nuclear stain DAPI (Molecular Probes) and mounted onto slides using AquaMount (Thermo Scientific). Immunofluorescent images were captured using a Zeiss LSM780 laser scanning confocal microscope.

Bi-Fluorescence Complementation (BiFC). To validate protein interactions, BiFC is based on the association of fluorescent protein fragments that are attached to components of the same macromolecular complex. 293 HEK cells plated on fibronectin-coated coverslips were transfected with expression plasmids using Lipofectamine 2000 reagent (Invitrogen) 24 h after plating and coverslips were processed 24 h after transfection. Coverslips were washed in PBS, fixed in 4% paraformaldehyde. Following three washes, cells were stained with the nuclear stain

59 DAPI (Molecular Probes) and mounted onto slides using AquaMount (Thermo Scientific). Immunofluorescent images were captured using a Zeiss LSM780 laser scanning confocal microscope.

Pull-down for active small Rho GTPases (Rac and Cdc42). To create GST-CRIB (Pak) pGEX2T-CRIB fusion proteins were induced in BL21 Escherichia coli by the addition of 1 mM isopropyl-1-thio-β-D-galactopyranoside for 3 h at 37 °C. After lysis and sonication, GST fusion protein was coupled to glutathione-agarose beads (Sigma), for 30 min at 4 °C. Protein concentration was estimated by resolving aliquots on SDS- polyacrylamide gels and staining with Coomassie Blue or run on TGX Stain-Free™ FastCast™ acrylamide gels (BioRad) and activated and imaged for total protein content as described above. An in vitro binding assay was performed by incubating lysates of transiently transfected 293 cells with 20 uL of GST-CRIB beads for 1 h at 4 °C, in the presence of 1 mM dithiothreitol. After three washes, proteins were eluted in 1X Laemmli buffer, boiled for 5 minutes at 105 °C, and loaded onto SDS- polyacrylamide gels. Immunoblots were carried out as described above.

Rac1 assay based on bioluminescence resonance energy transfer (BRET). 293 HEK cells were transiently transfected in complete media in 60 mm plates, with the various expression plasmids in addition to the RlucII-CRIB and rGFP-CAAX (prenylated rGFP) BRET biosensors, generously donated by Dr. Stephane A. Laporte. 24 h post-transfection cells were trypsinized, counted and seeded onto poly-ornithine-coated 96-well white plates, at a density of 25,000 cells per well, in triplicate for each condition and grown overnight in complete media. The next day, the media was replaced with 80 uL of Tyrode’s buffer (140 mM NaCl, 2.7 mM KCl, 1 mM

CaCl2, 12 mM NaHCO3, 5.6 mM D-glucose, 0.5 mM MgCl2, 0.37 mM NaH2PO4, 25 mM HEPES, pH 7.4) with 5 uM colenterazine 400a as substrate. Cells were left unstimulated or stimulated for 5 minutes with 100 ng/ml EGF. For kinetic measurements, BRET signals were read at 1, 3, 5, 7 and 9 minutes, using a Synergy2 (BioTekÒ) microplate reader. Filter set was 410 ± 80 nm and 515 ± 30 nm for detecting the RlucII Renilla luciferase (donor) and GFP10 (acceptor) light emission. BRET ratio was determined by calculating the ratio of the light emitted by GFP10 over the light emitted by the RlucII. Raw BRET was calculated as the ratio of GFP10

60 over RlucII and subtracted from unstimulated BRET and relative BRET was plotted on a graph as the difference in stimulated BRET minus unstimulated BRET. The specificity of the Rac1 signal was validated by inhibition with dominant negative Rac1 N17.

61 RESEARCH FINDINGS

In order to identify novel partners of Dok-4 Dr. Lemay's laboratory had previously performed yeast two-hybrid (Y2H) screens in a mouse kidney cDNA library and identified SHIP1 and Elk-4 [149], [174]. Among the candidate partners, another molecule was of particular interest, the Rac1 GTPase activating (GAP) protein β2-chimaerin-2 (Chn2). Two fragments of Chn2 were recovered in the screen performed in the presence of a co-expressed Src kinase. The shortest of the Chn2 fragments comprised codons for amino acids 1 to 182 (Figure 1A). This fragment contained the entire SH2 domain of Chn2. In addition, upon close inspection of the tyrosine residues of this fragment, we noticed that one of them, tyrosine 153 (Y153), was contained within the sequence NPIY, a canonical PTB domain-binding motif (NPXY). Interestingly, the public database PhosphositePlus contained proteomics data suggesting that Y153 is frequently phosphorylated (data not shown). In addition to its SH2 domain and putative PTB-binding motif, full-length Chn2 contains a diacylglycerol-binding C1 domain and a Rac1- specific GAP domain (Figure 1B).

To validate the interaction of Dok-4 and Chn2, we performed co-IP experiments in 293HEK cells transfected with an EGFP-tagged Chn2 N-terminal fragment and Myc-tagged Dok-4 with and without co-expression of an activated form of the Src family kinase Fyn (Fyn Y528F) (Figure 1C). This confirmed the interaction of Dok-4 with Chn2 and its enhancement by co-expression of a tyrosine kinase. In addition, we found that the intact Dok-4 PTB domain (aa 100-246) was necessary and sufficient for the interaction with Chn2, as it is interrupted by mutation of the Dok-4 PTB domain at a highly conserved and critical glycine residue (G207).

62 A C NPIY153 1 SH2 182

PLCɣ

TKs DAG

NPIY153

1 SH2 C1 RacGAP 468

Rac1

Figure 1: Interaction of Dok-4 with the Rac1 GAP β2-chimaerin-2 (Chn2). A: In a yeast two- hybrid screen for Dok-4 partners, two fragments of Chn2 were recovered in the presence but not in theFigure absence 1: Interaction of a co- expressedof Dok-4 with Src the kinase. Rac1 GAPThe shortestβ2-chimaerin Chn2-2 (Chn2).fragment A: isIn representeda yeast two-hybrid here. It containsscreen a for known Dok-4 phosphorylationpartners, two fragments that, of which Chn2 were we recoveredrecognized in the as presence being part but not of in canonical the absence PTB of a co-expressed Src kinase. The shortest Chn2 fragment is represented here. It contains a known domainphosphorylation-binding motif that, (NPXY). which we recognizedB: Overall as structure being part of of Chn2, canonical including PTB domain the diacylglycerol-binding motif -binding C1 domain,(NPXY). B the: Overall GAP structure domain of and Chn2, the including putative the PTB diacylglycerol-binding - motifbinding containing C1 domain, tyrosinethe GAP 153. C: domain and the putative PTB-binding motif containing tyrosine 153. B: NPxY153 and the surrounding Usingsequence co-IP ofis highly Myc taggedconserved Dok between-4 and mouse EGFP and-tagged human Chn2 Chn2 N and-terminal Chn1. C fragment,: Using co- IPwe of confirmed Myc tagged the interactionDok-4 and of EGFPDok--4tagged with Chn2Chn2 N and-terminal its enhancement fragment, we confirmedby co-expression the interaction of a oftyrosine Dok-4 with kinase Chn2 (here and its enhancement by co-expression of a tyrosine kinase (here constitutively active Fyn) (lane 3 vs 2). constitutivelyIn addition, activewe show Fyn) that the(lane intact 3 vs Dok 2).-4 InPTB addition, domain (aawe 100 show-246) that is necessary the intact and Dok sufficient-4 PTB for domain the (aa interaction100-246) with is necessary Chn2 (lanes and 5 vs sufficient 3), as it is interrupted for the interaction by mutation with of the Chn2 Dok- 4 (lanes PTB domain 5 vs at 3), a as it is critical glycine residue (lanes 7 vs 5). interrupted by mutation of the Dok-4 PTB domain at a critical glycine residue (lanes 7 vs 5).

To confirm the potential relevance of this novel interaction in renal IRI, we obtained lysates from kidneys of mice subjected to unilateral IRI and immunoblotted for Dok-4 and Chn2. While Dok-4 was detected in both sham and reperfused kidneys, Chn2 was only clearly detectable is reperfused kidneys after 24 hours of reperfusion (Figure 2).

Figure 2: Expression of Dok-4 and Chn2 in normal and reperfused kidney. In a mouse model of unilateral ischemia (I) and contralateral sham surgery (S), Chn2 protein expression was strongly upregulated after 24 hours of reperfusion.

To determine the structural basis of the Dok-4/Chn2 interaction, we used the Dok-4 PTB domain fused to glutathion S-transferase (GST) to perform pull-down assays from transfected 293HEK cell lysates. In the presence of activated Fyn, robust interaction of Dok-4 PTB with Chn2 was seen (Figure 3). However, when tyrosine 153 was mutated to phenylalanine, which has a similar structure to tyrosine, yet cannot be phosphorylated, the interaction was dramatically reduced. When an alanine mutation was introduced instead, with its structure differing from that of tyrosine and phenylalanine, the interaction was completely abolished.

Figure 3: The Dok-4 PTB domain interacts with phosphorylated tyrosine 153 of Chn2. A: GST or GST-Dok-4 PTB-coupled beads were used to pull down full-length Chn2 WT or Y153 mutants co-expressed with activated Fyn or empty vector. Interaction of Dok-4 and Chn2 was validated by pull-down of Chn2 with GST-Dok-4 PTB and shown to be dependent on co- expression of a tyrosine kinase (in this case Fyn) to phosphorylate Y153.

β2-chimaerin (Chn2) is highly homologous to another Rac1 GAP, α2-chimaerin (Chn1). Indeed, when we carefully inspected the sequence of Chn1, we found that it also contained a canonical NPXY motif around tyrosine 143 (NPIY). Moreover, the 10 amino acids flanking this motif are conserved between mouse and human Chn1 and Chn2 (Figure 4A), suggesting that Chn1 might also interact with Dok-4. To test this, we subjected Chn1 to pull-down assays and confirmed that it indeed interacted with the Dok-4 PTB domain in a tyrosine kinase-dependent manner (Figure 4B). Furthermore, this interaction was abolished by a tyrosine to alanine mutation at position 143 (Figure 4C). Thus, a tyrosine-based motif in Chn1 (Y143) and Chn2 (Y153) represents a previously unrecognized, yet highly conserved, PTB-binding motif that may regulate binding to Dok-4.

65 A B

153 145 AKMTTNPIYEHIGYm Chn2 145 SKMTTNPIYEHIGYhChn2 135 AKMTINPIYEHIGY mChn1 135 AKMTINPIYEHVGY hChn1

C D

Figure 4: Tyrosine 153 of Chn2 is contained within a highly conserved motif in human and murine Chn1 and Chn2. A: The NPxY153 motif and surrounding sequence is highly conserved between mouse and human Chn2 and Chn1. Y153 of Chn2 corresponds to Y143 of Chn1. B: As predicted. Dok-4 also interacts with Chn1 in the presence (lane 4) but not the absence (lane 2) of the co-expressed Src kinase Fyn. C: GST pull-down assay was performed on lysates of 293HEK cells transfected with Flag-tagged WT α2-chimaerin (Chn1) or its Y143A mutant in the presence or not of Src kinase (kinase/SH1 domain only, to avoid possible competitive binding of the Src SH2 domain). In addition to GST and GST-Dok-4 PTB, a GST-Vav2 SH2 domain was also tested for possible binding to Chn1, which turned out to be negligible. D: Following transient transfection in 293 cells, Dok-4 co-immunoprecipitates with full-length Chn1 in a manner that is enhanced by, but not entirely dependent on a co-expressed tyrosine kinase (here activated Fyn). Here, IP was performed with rabbit anti-Dok-4 antibody instead of monoclonal anti-Myc antibody, in order to minimize interference of mouse Ig heavy chains with the subsequent anti-Flag immunoblotting.

The Dok-4/Chn1 interaction of was also verified by co-IP (Figure 4D). However, repeated experiments showed that Dok-4 interacted much more readily with the isolated N-

66 terminal fragment of Chn2 than with full-length Chn2 (Figure 5). This was particularly apparent in co-localization studies, where expression of Dok-4 and Fyn caused a complete relocalization of the N-terminal fragment of Chn2 from a diffuse cytosolic pattern to one fully overlapping with the punctate membrane distribution of Dok-4 (Figure 6), whereas no convincing co- localization was observed with full-length Chn2 (data not shown). Taken together, this suggested that the conformation of full-length Chn2 may be unfavorable to the interaction with Dok-4.

Figure 5: Dok-4 interacts preferentially with the N-terminal fragment of Chn2 as compared to full-length Chn2. EGFP-tagged constructs of full-length (WT) Chn2 or N-terminal fragment (NT) Chn2 were transfected in 293 cells with or without Dok-4 and activated Fyn. Despite equivalent expression levels of WT and NT Chn2 in total lysates (left panel), co-IP with Dok-4 was much more prominent with truncated Chn2 as opposed to full-length Chn2, suggesting that WT conformation is unfavorable to the interaction.

67 A

Figure 6: The Chn2 N-terminal fragment co-localizes with Dok-4. Confocal microscopy was performed to visualize EGFP-tagged N-terminal Chn2, which contains Y153, and Myc-tagged Dok-4 (detected by anti-Myc primary antibody and Alexa555-coupled secondary antibody). A: In 293 HEK cells transfected with EGFP-Chn2 alone, diffuse cytoplasmic distribution is seen. B: In the presence of Dok-4 and activated Fyn, Chn2 NT rearranges into a distinct punctate distribution fully overlapping with that of Dok-4. We have previously defined this Dok-4 distribution as membrane-associated and PH domain-dependent [144]).

To clarify this and to more readily detect co-localization of Dok-4 with full-length Chn2 and to prove that it involved a very close interaction, we used bi-molecular fluorescence complementation (BiFC). This method is based on the observation that GFP derivatives can be split into N and C-terminal fragments that are non-fluorescent, but that close interactions can allow them to restore normal conformation and fluorescence (Figure 7A). Notably, this process involves protein folding that is largely irreversible and can therefore theoretically amplify interactions that are relatively transient, through a cumulative effect. To perform these experiments, we used the EGFP derivative Venus, with its N-terminal fragment fused to Chn2 and its C-terminal fragment fused to Dok-4 (Figure 7B). V1-Chn2 and Dok-4-V2 were transfected in 293HEK cells together with active Fyn and empty mCherry vector (as marker of transfection efficiency) and assessed under epifluorescence microscopy. Very little interaction was seen between Dok-4 and Chn2 WT (Figure 7C). However, a Chn2 mutant known to adopt

68 an active (open) conformation (I130A) [222] yielded a strong interaction signal (green) (Figure 7D). To demonstrate the specificity of this signal, we used Dok-4 construct with a mutated PTB domain (G207A) and found that the interaction signal was greatly reduced (Figure 7E). This suggested that the Dok-4/Chn2 interaction might be regulated not only by tyrosine kinase activity, but also by other factors known to promote its active conformation, such as binding to diacylglycerol (through the C1 domain) or to active Rac1 (through the Rac1 domain).

69 A B Venus-N- Chn2 NPIY Venus-N SH2 C1 RhoGAP

Dok-4- Venus-C PH PTB Venus-C

Figure 7: PTB-dependent interaction of Dok-4 with full-length Chn2 is facilitated by the active conformation of Chn2. Because, in contrast to Chn2 NT, full-length Chn2 appeared to interact and co-localize poorly with Dok-4, bi-molecular fluorescence complementation (BiFC) was explored as an alternative strategy to demonstrate interaction. A: Schematic representation of

70 the principles of BiFC to assess molecular interactions. A fluorescent protein (in this case the GFP variant, Venus) can be split into an N-terminal and C-terminal portion, each separately fused to candidate interactors. Molecular interaction allows reconstitution of fluorescent response, whereas this is absent from fragments that fail come into close proximity. B: Representation of the constructs used. The Venus1 (NT) fragment was fused to the N-terminus of Chn2 while the Venus2 (CT) fragment was fused to the C-terminus of Dok-4. C: V1-Chn2 and Dok-4-V2 were transfected in 293HEK cells together with active Fyn and empty mCherry vector (as marker of transfection efficiency) and assessed under epifluorescence microscopy. Very little green signal is seen, indicative of poor interaction. D: When WT V1-Chn2 is replaced by a mutant with active conformation (I130A), interaction (green signal) with Dok-4 is very prominent. E: BiFC signal is greatly reduced when the Dok-4 PTB domain is mutated (G207A).

In order to gain insight into the functional impact of Dok-4/Chn2 interaction, we examined the impact of both molecules on EGF-induced Rac1 activity. However, this proved to be extremely difficult and unreliable using a traditional GST pull-down approach (data not shown), although we could detect activation by overexpression of the Rac1 GEF domain of Dock180 (Figure 8A), more robust but static stimulus. To circumvent the challenge of traditional Rac1 assays, we sought the help of Dr. Stéphane Laporte to develop a Rac1 assay based on bioluminescence resonance energy transfer (BRET). The principles of this assay are described in Figure 8B. Like fluorescence resonance energy transfer (FRET), BRET generates a fluorescence signal in an interaction-dependant manner. However, because the initial energy is derived from luminescence, it requires only addition of a substrate instead of laser excitation and it can therefore be performed more quickly and quantitatively on whole cell populations, in a 96-well plate format. Initial validation of the technique revealed time-dependent activation BRET signal in response to angiotensin II and EGF, which was blocked by a dominant negative Rac1 (Figure 8C). Subsequent experiments showed that Chn2 and Dok-4 cooperated to inhibit EGF induced Rac1 signal (Figure 8D). Finally, Dok-4 containing a deletion of the PH domain lacked any ability to enhance Chn2 inhibitory action, but this was potently restored by the addition of a membrane-targeting myristoylation signal (Myr-Dok-4 DPH) (Figure 8E).

71 A B

C D

72 Figure 8: Measuring the impact of Dok-4 and Chn2 on Rac1 activity. A: Using a traditional GST-Pax CRIB pull-down assay, we were unable to consistently detect EGF-induced Rac1 activity in 293 cells (not shown), but we could induce Rac1 activation by overexpressing the catalytic domain (DHR2) of the Rac GEF Dock180. B: In order to detect Rac1 with higher sensitivity and precision, we used a BRET biosensor system designed by our collaborator, Dr. Stéphane Laporte. The assay uses RLuc-CRIB as donor and membrane-targeted rGFP-CAAX as acceptor. Upon incubation with coelenterazine, RLuc-CRIB emits light at 410 nm. If membrane Rac1 is inactive (GDP-bound), rGFP is not excited. If membraned Rac1 is active (GTP-bound), RLuc/rGFP interaction is facilitated and rGFP becomes excited by RLuc emission and it emits itself at 515 nm. C: 293HEK cells were transfected with receptors and stimulated with ligands as indicated. Both angiotensin II and EGF dynamically enhance Rac1 BRET signal. In cells expressing dominant negative (DN) Rac1, EGF-induced Rac1 BRET signal is greatly attenuated. D: Dok-4 and Chn2 cooperate to inhibit EGF-induced Rac1 activity. This represents the average of 5 independent experiments +/- SEM. E: Similar experiments were carried out in 293 cells, this time using a Dok-4 DPH construct lacking the PH domain and therefore unable to localize at the cell membrane. This construct was unable to potentiate Chn2-mediated inhibition of Rac1. However, when a membrane-targeting myristoylation signal was added to the DPH construct (Dok-4 Myr- DPH), inhibition of Rac1 was rescued.

CHAPTER 3: DISCUSSION, FUTURE STUDIES AND CONCLUSIONS

74 DISCUSSION AND FUTURE STUDIES

The epidermal growth factor receptor, a versatile signal transducer, is an important cell surface receptor, regulating signaling pathways and cellular processes that mediate metabolism, transcription, cell-cycle progression, differentiation, development, cytoskeleton arrangement and cell movement, apoptosis and intercellular communication [1]. The involvement of the EGFR in multiple cellular events emphasizes the importance of regulating EGFR signal transduction. In fact, aberrant EGFR signaling is characteristic in the development and progression of many human cancers [20] and proper EGFR signaling underlies many reparative processes following renal IRI [10]–[14]. Despite a comprehensive understanding of the events leading to EGFR activation, less is known about the events that negatively regulate EGFR signaling. The focus of this study was to identify negative regulators of EGFR in renal epithelial cells. Dr. Lemay's laboratory had previously determined that Dok-4 could inhibit EGF-induced signaling such as activation of ERK [131]. However, the basis of this Dok-4 action remains unclear, particularly as, in contrast to Ret, EGFR cannot phosphorylate Dok-4 (data not shown) and cannot directly bind Dok-4 [143]. Interestingly, recent proteomics data obtained in Dr. Lemay's laboratory indicate that Dok-4 does indeed interact with EGFR in HeLa cells, though presumably through an indirect mechanism (data not shown). This strengthens the relevance of Dok-4 to EGFR signaling. We have now identified a novel component of this EGFR/Dok-4 signaling pathway: the RacGAPs α2-chimaerin and β2-chimaerin, which are direct binding partners of Dok-4. We have demonstrated that Dok-4 and β2-chimaerin, act as regulatory factors, inhibiting Rac activation downstream of the EGFR in renal epithelial cells.

As an adaptor protein, the function of Dok-4 presumably relies on interactions with binding partners mediated largely by its PTB domain and C-terminal region. Therefore, in order to identify possible binding partners, we performed a yeast-two hybrid screen of a mouse kidney cDNA library, using full-length mouse Dok-4 as bait. Two screens were performed with and without the co-expression of the Src kinase, Lyn, to determine kinase-dependent and independent interactions. From these screens we identified the RacGAP, β2-chimaerin, as a kinase-dependent Dok-4 interacting partner, containing a canonical PTB binding NPIY motif (Figure 1A and 1B).

75 The Dok-4 PTB domain has been historically defined as an IRS-like PTB domain, based on its sequence homology to IRS1 [111]. We had previously redefined the borders of the Dok-4 PTB domain to include an extended C-terminal alpha-helix [174], and using these redefined borders, we investigated the kinase-dependent interaction between Dok-4 PTB domain and the RacGAPs, β2-chimaerin. Importantly, through co-immunoprecipitation, we confirmed that Dok- 4 interacts readily with the β2-chimaerin N-terminal (NT) fragment, that the Dok-4 PTB domain is sufficient for this interaction and that the binding of the β2-chimaerin NT to Dok-4 is enhanced by co-expression of a tyrosine kinase. Additionally, a G207A mutation, disrupting the structure of the Dok-4 PTB domain alters the ability of Dok-4 to bind β2-chimaerin (Figure 1C), further confirming the role of the Dok-4 PTB domain in this interaction.

We looked at expression levels of Dok-4 and β2-chimaerin in mice, during the recovery phase following renal IRI and found both to be upregulated in the injured kidney (Figure 2). There is, however, a 24 hour difference in the time of upregulation of Dok-4, which peaks 30 minutes post IRI, versus β2-chimaerin, which peaks 24 hours post IRI. This difference may reflect the physiological roles of these proteins in IRI, where immediately following initial injury, Dok-4 may need to be uncoupled from β2-chimaerin, with the interaction of these two proteins becoming relevant in the recovery phase, post-IRI.

We determined that the interaction between the Dok-4 PTB domain and β2-chimaerin is dependent on tyrosine phosphorylation of a canonical NPIpY PTB-binding motif, located in the N-terminal SH2-C1-linker region of β2-chimaerin. Mutation of the chimaerin NPIY motif to NPIF or NPIA effectively inhibited the interaction with Dok-4 (Figure 3), suggesting that specific phosphorylation of the PTB-binding NPIY motif is necessary for the interaction and defining the interaction between Dok-4 and β2-chimaerin as a “typical” PTB interaction.

We later identified the homologous α2-chimaerin isoform, containing a similar NPIY motif (Figure 4A) as a potential Dok-4 binding partner. As predicted the Dok-4 PTB domain was sufficient to interact with α2-chimaerin and did so in a kinase-dependent manner (Figure 4B); and like β2-chimaerin, phosphorylation of the α2-chimaerin PTB-binding NPIY motif is necessary for the interaction with Dok-4 (Figure 4C), defining the interaction as a “typical” PTB

76 interaction; however, Dok-4 co-immunoprecipitates with full-length Chn1 in a manner that is enhanced by, but not entirely dependent on a co-expressed tyrosine kinase (Figure 4D), suggesting additional binding regions, such as the Dok-4 C-terminal region may participate in the interaction between full-length Dok-4 and α2-chimaerin. Additional studies with various Dok-4 mutants, lacking the PTB domain, will help identify additional binding sites.

Since binding of Dok-4 with the transcription factor, Elk-4, has been identified as occurring through an “atypical” PTB domain interaction [149], likely distinct from the canonical PTB fold, we would be interested in performing GST-PD and Co-IP assays to determine if Elk-4 and β2-chimaerin or Elk-4 and α2-chimaerin can simultaneous bind Dok-4, and further qPCR, cell proliferation and Rac activation assays could tell us the functional effect of simultaneous binding on cellular proliferation and Rac activation.

We suspect that the conformation of full-length β2-chimaerin hinders binding to Dok-4, given that the β2-chimaerin NT co-immunoprecipitates with Dok-4 more robustly than the full- length protein, despite equivalent expression levels (Figure 5), suggesting that, in addition to tyrosine kinase, other stimuli, such as EGFR activation, or co-factors, such as Nck1 or serine/threonine kinases may be necessary to enhance binding between full-length proteins. This finding corroborates with prior studies, showing that under basal conditions, β2-chimaerin is autoinhibited in the cytosol [196] (Figure J). We did see co-localization between over-expressed Dok-4 and the β2-chimaerin NT, in the presence of tyrosine kinase (Figure 6B); however, full- length β2-chimaerin failed to co-localize with Dok-4, when over-expressed with tyrosine kinase (Figure 7C). Dr. Marcelo Kazanietz’s group had identified an Ile to Ala mutation in position 130 of β2-chimaerin, a residue in the SH2 domain that makes contact with the C1 domain, essentially mimicking the active conformation, greatly increasing both the sensitivity to PMA-induced translocation as well as Rac-GAP activity in vitro and hypothesized that this mutant adopts a conformation that is energetically favorable for ligand binding [222]. Indeed, we found the active (open) β2-chimaerin I130A mutant to co-localize with Dok-4, when over-expressed with tyrosine kinase (Figure 7D), and the interaction was abolished by the Dok-4 G207A mutation, which disrupts the PTB domain structure (Figure 7E), suggesting that β2-chimaerin activation requires a major conformational change in order to expose the canonical PTB-binding motif. In order to

77 further elucidate the mechanism of interaction between Dok-4 and full-length β2-chimaerin, future studies can look into the ability of β2-chimaerin to bind Dok-4, as well as the phosphorylation state of β2-chimaerin, specifically Y153, in response to EGFR activation, which is known to induce an open-active β2-chimaerin conformation [194].

Looking into the functional effect of the interaction between Dok-4 and β2-chimaerinon membrane-localized Rac1 activity, downstream of EGFR activation, we utilized a highly sensitive BRET biosensor system (Figure 8B) designed by our collaborator, Dr. Stéphane Laporte. We first validated the ability of the system to register EGF-induced Rac1 activation, using angiotensin II and its receptor, which are known to activate Rac1 [223], as a positive control and dominant negative Rac1 as a negative control (Figure 8C). We found that in 293 HEK cells, which do not endogenously express Dok-4 or β2-chimaerin, co-transfection of Dok-4 and β2-chimaerin had a potent inhibitory effect on EGF-induced Rac1 activation. This effect was not seen when the cells were transfected with either Dok-4 or β2-chimaerin alone, (Figure 8D) suggesting that Dok-4/β2-chimaerin interaction results in cooperative inhibition of Rac1. Interestingly, the Dok-4 ΔPH construct, lacking the PH domain and therefore unable to localize at the cell membrane, was unable to potentiate β2-chimaerin-mediated inhibition of Rac1; however, when a membrane-targeting myristoylation signal was added to the ΔPH construct (Dok-4 Myr-ΔPH), inhibition of Rac1 was rescued (Figure 8E). These results suggest that Dok-4 functions to recruit active β2-chimaerin to the cell membrane, where it can exert its RacGAP activity on active Rac1. More experiments are needed to clarify the contribution of Cdc42 vs Rac1 to the BRET signal, and the assays must be repeated to investigate whether co-expressing Dok-4 and α2-chimaerin has a similar inhibitory effect on Rac1 activity.

The downstream effects of Rac1 modulation by the interaction between Dok-4 and β2- chimaerin have yet to be identified. We have previously observed that Dok-4 knockdown enhances EGF-induced cellular proliferation and have determined that Dok-4 inhibits the immediate early genes Egr-1 and Fos as well as cyclin D1, negatively regulating cell cycle progression and proliferation through direct binding of the transcription factor Elk-4, leading to its destabilization [149] (Figure 9A). Based on the current work, we postulate that downstream of RTK activation, in addition to its interaction with Elk-4, Dok-4 exerts its anti-proliferative effect

78 through the recruitment of the RacGAP β2-chimaerin, effectively inhibiting cyclin D1 through inhibition of the Rac1 pathway (Figure 9A). Notably, it has already been shown that following RTK activation, auto-inhibited cytosolic β2-chimaerin (Figure 9B) undergoes a conformational change and translocates to the membrane via the generation of membranous diacylglycerol by PLCγ and recruitment of the β2-chimaerin C1 domain, promoting and maintaining β2- chimaerin’s open, active conformation (Figure 9C). We propose an alternative model by which Dok-4 may facilitate β2-chimaerin activation and membrane recruitment through the binding of the Dok-4 PTB domain to tyrosine phosphorylated β2-chimaerin, such that the Dok-4 PTB domain maintains β2-chimaerin in its active conformation, while the Dok-4 PH domain binds

membrane phospholipids (Figure 9D).

C1 A B SH2

RacGAP

C RTK

PIP2 DAG

P P PLCɣ

NPIY153 SH2 C1 RacGAP

Rac1 D RTK

PIP2 PH P P PLCɣ PTB NPIpY153 SH2 C1 RacGAP

RTK/SFK Rac1

Figure 9: Model of Dok-4 anti-proliferative action. A: We propose that Dok-4 inhibits cyclin Figure 9: Model of Dok-4 anti-proliferative action. A: We propose that Dok-4 inhibits cyclin D1 D1expression expressionand andcell cellproliferation proliferationthrough throughinhibition inhibitionof the Rac of1 thepathway Rac1 throughpathwayrecruitment through recruitmentof the Rac1 GAP Chn2. Dok-4 also inhibits cyclin D1 induction by binding and inhibiting the transcription factor Elk-4. B-D: Proposed model of how Dok-4 may facilitate recruitment of Chn2 to the membrane. In the basal state, Chn2 exists in an inactive, closed conformation (B). Following RTK 79 activation, diacylglycerol (DAG) is produced in the cell membrane by the action of PLCγ. DAG interacts with the C1 domain, resulting in activated (open) Chn2 that translocates to the membrane (C). In the presence of Dok-4, an alternative model of Chn2 activation and membrane recruitment is possible through binding of the Dok-4 PTB domain to tyrosine phosphorylated Chn2. The Dok-4 PTB domain maintains Chn2 in an open conformation while the Dok-4 PH domain binds membrane phospholipids such as phosphatidylinositol 4,5 bisphosphate (PIP2). of the Rac1 GAP Chn2. Dok-4 also inhibits cyclin D1 induction by binding and inhibiting the transcription factor Elk-4. B-D: Proposed model of how Dok-4 may facilitate recruitment of Chn2 to the membrane. In the basal state, Chn2 exists in an inactive, closed conformation (B). Following RTK activation, diacylglycerol (DAG) is produced in the cell membrane by the action of PLCγ. DAG interacts with the C1 domain, resulting in activated (open) Chn2 that translocates to the membrane (C). In the presence of Dok-4, an alternative model of Chn2 activation and membrane recruitment is possible through binding of the Dok-4 PTB domain to tyrosine phosphorylated Chn2. The Dok-4 PTB domain maintains Chn2 in an open conformation while the Dok-4 PH domain binds membrane phospholipids such as phosphatidylinositol 4,5 bisphosphate (PIP2).

Since Dok-4 has been implicated in the negative regulation of mitogenic signaling, it is possible that these proteins function to limit cellular proliferation during the recovery phase following IRI, when surviving tubular epithelial cells are rapidly proliferating, thereby inhibiting the development of renal fibrosis. We will look into the expression level of renal α2-chimaerin following IRI. Additionally, it will be important to address the potential role of each protein as well as the role of the interaction between Dok-4 and α2- and β2-chimaerin in vivo, using knockout mice. A β2-chimaerin knockout mouse [224] and an α2-chimaerin knockout mouse [197] already exist, and show a minimal phenotype, but seeing as how α2-chimaerin appears to function in much the same way as β2-chimaerin, we wonder about the effect of a combined α2- chimaerin/β2-chimaerin knockout phenotype, especially under injury challenge such as renal IRI. The specific role of the interaction between chimaerins and Dok-4 could be studied in mice by introducing a point mutation in α2-chimaerin Y143/β2-chimaerin Y153 with CRISPR/CAS9. We will also test the impact of Dok-4 knockout on recovery and cellular proliferation following renal IRI. We have generated a unique global Dok-4 knockout mouse, which is viable and shows no apparent phenotype until 9 months of age. As part of phenotypic characterization studies, we should subject these mice to the challenge or renal IRI to see if it displays enhanced proliferation and/or recovery following renal injury.

CONCLUSIONS

Collectively, the work presented in this thesis identifies two novel binding partners of the adaptor protein Dok-4. We found that the major binding site between Dok-4 and α2-chimaerin and Dok-4 and β2-chimaerin involves the Dok-4 PTB domain and Y143 of α2-chimaerin and Y153 of α2-chimaerin and that the interactions with Dok-4 are enhanced by tyrosine phosphorylation at these positions. Functionally, Dok-4 acts to recruit β2-chimaerin to the membrane where they cooperate to inhibit EGFR-induced Rac1 activity. Though it remains to be investigated, we suggest that the inhibition of Rac1 following renal IRI will limit renal repair. Furthermore, we speculate that the mechanism of Rac1 regulation proposed in this thesis will be applicable not only following renal IRI, but also during oncogenesis where Rac activity is often increased [208]. Interventions targeting Dok-4 in the negative regulation of mitogenic signaling, through the inhibition of Dok-4 expression or interference with the recruitment and activation of chimaerins may enhance renal recovery following IRI, while interventions enhancing the recruitment and activation of chimaerins by Dok-4 may limit the progression of cancer [225] and improve patient outcome.

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